Previous Article | Next Article 
Microbiology and Molecular Biology Reviews, September 2005, p. 440-461, Vol. 69, No. 3
1092-2172/05/$08.00+0 doi:10.1128/MMBR.69.3.440-461.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
Fate of Heterotrophic Microbes in Pelagic Habitats: Focus on Populations
Max Planck Institute for Marine Microbiology, Bremen, Germany
SUMMARY INTRODUCTION Units of Interest: Single Microbial Populations Marine Autotrophic Picoplankton as a Model Microbial Phenotypes Operational Phylogeny-Based Definition of Microbial Populations Synoptic Discussion of Marine and Freshwater Assemblages TOOLS, TECHNIQUES, AND STUDY CONCEPTS 16S rRNA Gene Clone Libraries: Essential but Prone to Bias Single-Cell Identification Problems with FISH and Possible Solutions Quantification of Abundance and Biomass Single-Cell Activities of Individual Populations Microautoradiography. Fluorescent activity tracers and flow sorting Distribution and Dynamics of Different Populations Experimental Enrichments in the Field Defined Laboratory Communities and Pure Culture Studies OCCURRENCE OF MICROBIAL POPULATIONS IN PLANKTON A Few Words on Diversity Bacterial Populations in the Marine Water Column Marine Archaea Numerically Important Bacteria in Freshwater Bacterial Populations Attached to Organic Particles FACTORS CONTROLLING MICROBIAL POPULATION SIZES Competition for Different Substrates Patchiness and Gradients Mortality Viral lysis. Selective predation. OUTLOOK ACKNOWLEDGMENTS REFERENCES
 Top NextReferences |
|---|
Major biogeochemical processes in the water columns of lakes and oceans are related to the activities of heterotrophic microbes, e.g., the mineralization of organic carbon from photosynthesis and allochthonous influx or its transport to the higher trophic levels. During the last 15 years, cultivation-independent molecular techniques have substantially contributed to our understanding of the diversity of the microbial communities in different aquatic systems. In parallel, the complexity of aquatic habitats at a microscale has inspired research on the ecophysiological properties of uncultured microorganisms that thrive in a continuum of dissolved to particulate organic matter. One possibility to link these two aspects is to adopt a "Gleasonian" perspective, i.e., to study aquatic microbial assemblages in situ at the population level rather than looking at microbial community structure, diversity, or function as a whole. This review compiles current knowledge about the role and fate of different populations of heterotrophic picoplankton in marine and inland waters. Specifically, we focus on a growing suite of techniques that link the analysis of bacterial identity with growth, morphology, and various physiological activities at the level of single cells. An overview is given of the potential and limitations of methodological approaches, and factors that might control the population sizes of different microbes in pelagic habitats are discussed.
|
TopPreviousNextReferences |
|---|
Molecular biological methods centered on the rRNA gene have developed into powerful tools for the cultivation-independent identification of aquatic microorganisms. Over the last decades, the composition and diversity of microbial assemblages in numerous marine and freshwater environments have been studied by 16S rRNA gene cloning and sequencing (55, 76, 85, 91, 121, 303), community fingerprinting (36, 49, 156, 185, 254), hybridizations with oligo- or polynucleotide probes (2, 89, 136, 188, 218), or by combinations of these approaches (63, 144, 223, 266, 278).
A major goal of aquatic microbial ecology is to understand the specific roles of different microorganisms as mediators of element fluxes, e.g., during the remineralization of nutrients and organic carbon. So far, the novel molecular approaches have focused mainly on the occurrence and evolutionary relatedness of different bacteria, archaea, and picoeukaryotes. Clearly, there is a need to progress beyond a mere descriptive analysis of microbial diversity and community structure, to provide information about which microbes are involved in various biogeochemical processes. An integration of the more diversity-centered and the more biogeochemical perspectives in microbial ecology has been sought in the concept of the "structure and function" of microbial assemblages (73, 256). Such terminology might be fully appropriate for the study of technologically designed environments that serve particular functions, e.g., wastewater treatment plants, of coherent physiological groups, or of habitats like sediments or biofilms that feature a rigid physical structure (37, 257, 308). However, the concept of "structure and function" appears to overemphasize the "bottom-up" aspect of aquatic microbial assemblages, i.e., aspects that are related to microbial substrate and nutrient utilization. Pelagic habitats are also zones of high microbial mortality, and a substantial fraction of bacterial biomass may be transferred to higher trophic levels by predation (272). In order to understand the different heterotrophic microbes that inhabit the marine and freshwater plankton, it might thus be more adaeqate to consider both the specific "role" and the "fate" of microbial populations.
In many respects "aquatic microbial communities" are theoretical concepts rather than biologically real entities with (physical) structure, common genome or evolutionary history (66). Although ultimately there might be community assemblage rules also for the aquatic microbes (118), bacterioplankton assemblages could likewise be regarded as more or less loose collections of individual genotypic populations that change over time in features such as growth, mortality, and size. Here we discuss the advantages of adapting such a "Gleasonian" viewpoint (87), i.e., we focus on distinct bacterial populations that can be reliably identified and quantified in aquatic assemblages rather than on microbial community structure, diversity, or function as a whole. Specifically, we review current methodological approaches, and we assemble information about the occurrence, the phenotypic properties, and the possible role and fate of such defined microbial populations in the water column.
The advantages of a focus on single microbial populations can be illustrated by looking at autotrophic marine picoplankton. Prochlorococcus is an ubiquitous free-living phototrophic cyanobacterial genus that is common in temperate to tropical marine waters (205), where it may account for a high fraction of total primary production. This group was only discovered in 1989 by means of a then-novel technology, flow cytometry (38). The ability to reliably distinguish Prochlorococcus populations from other species of autotrophic marine picoplankton (e.g., Synechococcus) has resulted in concerted research about the ecology, physiology, and genomic constitution of these organisms that is unparalleled in aquatic microbial ecology. During the past 15 years, numerous strains of Prochlorococcus from various locations have been isolated and studied in the laboratory (205). The geographic distribution limits and the contrasting vertical niches of physiologically distinct genotypes have been established (67, 205, 314). Recently, the capacity of Prochlorococcus to heterotrophically utilize organic nitrogen sources has been described both in situ (327) and as a feature of its genome (246). The successful unraveling of the functional role of this particular bacterial group based on its reliable in situ identification might also provide a conceptual model for the study of heterotrophic microbes in the water column.
Major biogeochemical processes in the water column are related to the activities of heterotrophic microbes, e.g., the mineralization of organic carbon from photosynthesis or its transport to higher trophic levels (13). This is deduced from bulk measurements of microbial growth, respiration, the rates of biomass production, and its loss to predation or to viral lysis (53, 74, 77, 142, 270). Such bulk parameters sum up the potentially different rates of cell replication and mortality of various microbial species. While this may be essential to follow the fate of organic matter as a whole, it has also separated aquatic microbial ecology from other fields of microbiology that insist on well-identified microorganisms as the basic units of research. During the last decade the complexity of microbial phenotypes has moved into the focus of research that aims to explain biogeochemical processes. Our view of microbes as mediators of element flow has matured from the black-box approach of the early food web models to a subtler understanding of the ecophysiology of organisms that thrive in a heterogenous environment of dissolved and particulate organic matter (12).
The diversity of phenotypes within aquatic microbes reflects the complexity of the habitats at a microscale (3, 106, 206, 281). For example, a large fraction of cells in coastal bacterioplankton may be motile during particular seasons or times of the day, and the duration of their actual swimming periods may be affected by substrate availability and patchiness (103). This might be related to chemosensory functions that allow bacteria to actively accumulate in substrate-rich microniches such as the phycosphere of senescent algae (15, 23). The balance between particle colonization efficiencies and detachment dynamics may even reflect species-specific microbial life cycles (100, 140, 141). Culturable genera of aquatic bacteria harbor large numbers of facultative anaerobes, and ten percent of bacteria in oxic coastal marine waters could incorporate glucose both at oxic and at anoxic conditions (4, 243). Some strains isolated from the plankton are capable of degrading complex organic molecules or of utilizing organic sulfur compounds (94, 95). Other aquatic bacteria differ in the duration of the lag phase that precedes growth in batch culture (212), exhibit high tolerance to UV radiation (290), secrete or resist antibiotics (61, 161), or are capable of cell communication by quorum sensing (97). In addition, indications of microbial activity are found in the water column that presently cannot be assigned to particular bacterial groups, e.g., microbial exoenzymes with contrasting kinetic properties (9, 307).
Aquatic microbes have moreover developed a range of specific defense strategies. Some freshwater bacteria form threadlike morphotypes or microcolonies that secrete large amounts of exopolymerous substances, thereby obtaining protection against ingestion by bacterivorous protists (113, 223). In coastal North Sea surface waters >25% of prokaryotes were covered by a polysaccharidic capsule, a feature of many pathogenic species (288) which again might be related to grazing resistance. A high percentage of bacteria in different freshwaters are gram-positive (263), which enhances their protection against digestion by unicellular eukaryotes (114, 125).
However, one must not conceptually merge distinct microbial species into fictional chimeric "aquatic bacteria" (Fig. 1). More appropriately, the various phenotypic features should be regarded as individual elements of unique life strategies that explain the occurrence and distribution patterns of different genotypes in aquatic habitats. Some of these phenotypic features, such as motility or filament formation, may be widespread across very different phylogenetic lineages (115), whereas others, e.g., antibiotic production or resistance, may be highly characteristic for particular species or genera (100, 160). A single genotype may express a variety of phenotypic features or even phenotypic plasticity (90, 115), and identical features may be shared by numerous genotypes. Thus, a central task of microbial ecology is to assign the various physiological abilities and phenotypes observed in situ to particular microbial species or genotypes.
 View larger version (27K): [in a new window] |
FIG. 1. Schematic example of phenotypic diversification of aquatic bacteria. The relationship between the different genotypes and their specific ecophysiological features, life strategies, and behavior is often poorly understood.
|
It is still the subject of debate whether there can be a natural species concept in microbiology (41, 248). Therefore, a definition of "populations" in field investigations of microbial ecology should be more flexible than, e.g., for laboratory studies of ecophysiology or evolution (27, 64, 128). In our view, which is shaped by the available tools and the lack of information about the ecology of 16S rRNA-defined populations in the environment, almost any phylogenetically coherent group of microbes could potentially qualify as a "meaningful" population. The phylogenetic resolution that adequately delineates microbial populations largely depends on the ecological goals of a specific investigation. A meaningful population should represent a unit that (i) is recognizable in subsequent investigations and (ii) exhibits ecologically distinct features from other recognizable units (Fig. 2).
 View larger version (30K): [in a new window] |
FIG. 2. Conceptual model for an operational definition of microbial populations. In the context of ecological investigations three bacterial taxa, A, B, and C, can be meaningfully defined as different populations if they also differ in an ecologically relevant feature (parameter 1). However, a splitting of population C may be required if it is studied in the context of a second environmental variable (parameter 2).
|
The distinction of genotypic compartments within mixed microbial assemblages helps to compile ecologically relevant information about the abundances and activities of phylogenetically closely related groups of microorganisms, e.g., about their horizontal or vertical distribution patterns (210, 295), seasonality (63, 136), occurrence on particles (261), cell morphology (223), growth response to confinement (61, 72, 254), sensitivity to predation (16), incorporation of particular substrates (47, 169, 306), or growth under anoxic conditions (4). Our conceptual framework thus encompasses investigations of the seasonal occurrence of pathogenic Vibrio spp. in coastal waters (83, 117), of the response of a specific clade of freshwater betaproteobacteria to food web manipulations (278), but also of the differences in the vertical distribution of Bacteria and Archaea in the open ocean (136). On the other hand, it may also refer to the relative contributions of genotypic populations to a single biogeochemical process, e.g., the roles of planktonic members of the Roseobacter or SAR11 lineage in sulfur cycling (96, 169, 170, 306, 323).
It should be noted that a microbial population that is defined above the level of the single clone might represent an operational rather than a natural category. Therefore, such an operational taxonomic unit will be limited in its usefulness to a particular study context, and it should be refined or expanded if found inadequate for a specific purpose (Fig. 2). Some phylogenetic clades might harbor organisms with closely related or even identical 16S rRNA gene sequences but rather different ecophysiological properties (128). Sometimes it may be useful to merge such strains into larger categories, e.g., to study the occurrence of genera associated with particular phytoplankton blooms (96, 323). At other instances it might be necessary to distinguish between closely related strains, e.g., to understand the ecological significance of physiological variability (27, 109, 128) (Fig. 2).
A number of fundamental similarities in the functioning of marine and freshwater microbial assemblages have been described, in particular carbon transfer and food web structure (203, 251, 313) and the role of bacteria attached to aggregates (105, 229). This has traditionally led to a fruitful exchange between limnological and oceanographic research on planktonic microbes, based on shared concepts and on a common set of techniques (79, 81). Recently, the relationship between the two disciplines has somewhat deteriorated, conspicuously paralleled by a shift in research focus from food webs and element cycles to microbial biodiversity. In fact, cultivation-independent approaches have revealed by far more differences (89, 180) than similarities (14) in the composition of the microbial assemblages in the two habitat types.
Can identical ecological concepts be meaningfully applied to communities composed of bacterial groups from completely different phylogenetic lineages? Our review may in parts substantiate such doubts, e.g., with respect to the role of allochthonous (156) or filament-forming (130) microorganisms in the two habitat types. Yet at the same time we illustrate that a focus on individual microbial populations also provides a platform to compare the ecological value of particular phenotypic traits, e.g., of motility and small cell size (103, 221, 238), or of specific growth strategies (65, 68, 212) in very different genotypes and assemblages across aquatic habitats.
The scope of this review is limited in several dimensions. We do not attempt to draw an exhaustive picture of the microbial ecology of heterotrophic aquatic microbes. Instead we try to integrate a rather diverse range of facets, focusing on current methodological and experimental approaches, and on empirical findings related to traditional topics of population ecology. We argue that during the last decade the study of aquatic microbial populations has developed into a rewarding field of research that is based on mature methodology. Since our focus lies on the ecology of different populations rather than on the study of total assemblages, we only marginally touch the rich literature on genetic fingerprinting or on the diversity of aquatic communities. We limit our discussion to the "ordinary saprophytes", i.e., to the predominantly chemoorganoheterotrophic microbes (although admittedly the borders between purely heteroorganotrophic and photoautotrophic energy acquisition by pelagic microbes do become more and more fuzzy [20, 327]). We ignore microbial assemblages in sediments, biofilms, or in engineered systems, and we do not discuss in detail the physicochemical environments of aquatic microbes, but rather point to more specialized reviews, e.g., about macroscopic organic aggregates or exopolymerous substances (206, 281).
|
TopPreviousNextReferences |
|---|
During the last 15 years aquatic microbial ecology has been shaped by a diversification of methods for the cultivation-independent study of microbial identity, activity, and genomic constitution (11). This trend is reflected in the recent foundation of a scientific journal on methods in limnology and oceanography that is edited by a microbial ecologist (http://aslo.org/lomethods/editor.html). The following section does not give an exhaustive listing of tools for the study of microbial populations in situ. Instead, it covers a suite of complementary approaches which have been applied in various combinations to analyze bacteria in environmental samples. They can provide a coherent framework for ecological investigations about the abundant microbes in the water column, i.e., taxa that contribute approximately 1% or more to the total picoplankton assemblages. Other approaches, e.g., quantitative PCR (17, 283), might be more appropriate to study microbial populations that are substantially smaller.
|
TopPreviousNextReferences |
|---|
In order to define the important populations in a new environment or at a particular time point, it is often necessary to conduct a prior study about microbial diversity, most commonly of 16S rRNA genes. Full or partial sequences can be amplified from extracts of environmental DNA (75, 301) or directly from cells concentrated on membrane filters (21, 145) by PCR with appropriate primer sets (84, 194). These fragments are then ligated into plasmids, cloned into Escherichia coli, and the sequences of vector inserts from an appropriate number of clones are determined. The PCR step is omitted altogether in so-called shotgun clone libraries (305). Since such libraries require a drastically greater screening effort, they are used for obtaining metagenomic information rather than for the mere collection of 16S rRNA genes.
The diversity of sequences in PCR-generated clone libraries may often not quantitatively reflect the diversity of the sequence types that are present in the original sample (242, 294). Already the DNA extraction may introduce biases, e.g., against bacteria with gram-positive cell walls (69). Primers that are designed to target the majority of known bacterial 16S rRNA gene sequences may exhibit mismatches to unknown sequence types (45, 287), and the presence of particular sequence types in mixed DNA may influence the PCR amplification efficiencies of other templates (292).
As a consequence some phylogenetic groups of aquatic microorganisms are overrepresented in clone libraries, whereas others are absent. For example, bacteria affiliated to the Cytophaga/Flexibacter/Flavobacterium group of the Bacteroidetes were rarely found in 16S rRNA gene clone libraries from coastal marine water samples, yet they may represent one third of all bacteria in such habitats (47, 63). In a library of coastal North Sea surface waters 80% of sequence types were related to the marine SAR86 clade, whereas this group formed <10% of microbial cells (63). In addition, PCR may result in chimeric sequences from two templates, and it may thus even produce artificial sequence diversity (124, 155). These biases may affect not only clone libraries, but also PCR-based methods for the genotypic fingerprinting of microbial assemblages (183, 193).
Sometimes it is desirable to limit the analysis of microbial diversity to a defined phylogenetic subset, e.g., to marine archaea or to freshwater actinobacteria (173, 309). This can be achieved by PCR with primers that are specific for the 16S rRNA genes of the group of interest (287). Such primers may cause an underestimation of the potential diversity within a particular phylogenetic group, e.g., some uncultured freshwater actinobacteria show one or more mismatches with the available sets of specific primers (309). Moreover, sequences generated with specific primers often cover only a part of the total 16S rRNA gene, typically less than 1,000 base pairs (55, 287). Partial sequence information negatively affects the accuracy of phylogenetic reconstruction (163), and it limits the range of potential signatures for subsequent in situ population studies by hybridization. Preferably, clone libraries of almost complete 16S rRNA gene sequences should be produced with general bacterial or archaeal primers, and they should then be screened for particular groups of interest (173, 184, 309).
The potential target groups for population studies are defined from sequence data by reconstruction of phylogenetic relationships. This analysis aims at placing the new environmental sequence types into different clades or clusters and to establish stable branching patterns. It forms the base for a phylogenetically meaningful definition of single populations and for the design of the corresponding oligonucleotide probes. A discussion of the reconstruction of microbial phylogenies from 16S rRNA gene sequences would go beyond the range of this review, and we draw the reader's attention to specific publications on this subject (163, 164).
|
TopPreviousNextReferences |
|---|
Since 16S rRNA gene clone libraries do not accurately reflect the abundances of microbes from particular phylogenetic clades in the environment, such analysis needs to be complemented by other strategies for the study of single populations in situ. Hybridization techniques against extracted rRNA or intact cells have developed into an important tool of choice for this purpose. Our review is particularly focused on whole-cell fluorescence in situ hybridization (FISH) with rRNA-targeted oligo- or polynucleotide probes (6, 7, 58, 86, 211, 217). FISH allows the identification and quantification of different microbial taxa in environmental samples. It links information derived from molecular phylogeny with epifluorescence microscopy, a research tool that has considerably formed our understanding of water-column microbes (13, 122, 231).
The FISH analysis can be carried out at two levels (Fig. 3). For one, investigators may draw upon the rapidly expanding set of available probes (162) (Table 1). Most frequently, probes have been applied that are targeted to higher taxonomic levels, i.e., to Bacteria, Archaea, Eucarya, subgroups of Proteobacteria, gram-negative sulfate-reducing bacteria, clades of ammonium-oxidizing bacteria, etc. In an earlier review on FISH, such an analysis was proposed as a first step of a larger "top-to-bottom" strategy (7).
 View larger version (17K): [in a new window] |
FIG. 3. FISH, a versatile tool for the study of microbial populations in aquatic systems. The FISH approach is embedded in a suite of techniques that are required for the acquisition and processing of rRNA sequence information or that allow simultaneous investigation of microbial geno- and phenotypes at the single-cell level. S1 to S4, samples.
|
|
View this table: [in a new window] |
TABLE 1. Some oligonucleotide probes targeted to clades of marine and freshwater heterotrophic picoplankton
|
The second level of FISH analysis is more technically demanding. It includes the design and optimization of new FISH probes for individual phylogenetic clades or single phylotypes based on sequence information from 16S rRNA gene clone libraries of environmental DNA (Fig. 3). This strategy has been termed the "full-cycle rRNA approach" by one of the authors (7). The procedure is comparable to the testing of a scientific hypothesis. Specifically, it is verified if a particular clade of sequence types obtained from environmental DNA after PCR amplification indeed harbors microbes that form a substantial fraction of the assemblage in the studied habitat. The lower limits for a reliable quantification of cell numbers by the FISH approach as set by counting effort and negative controls typically range around 0.1 to 2% of total counts (217). Consequently, researchers applying FISH may need to test a number of newly designed specific probes on field samples to discover microbial populations that are sufficiently abundant for subsequent quantitative population studies. This implicit risk of falsification of a working hypothesis represents a major conceptual difference between the full-cycle rRNA approach and other strategies that analyze microbial assemblages by molecular fingerprinting of PCR-amplified rRNA genes (185, 254).
|
TopPreviousNextReferences |
|---|
In summary, FISH probes represent tools that are rather tedious to construct but easy to apply. The specificity of oligonucleotides for a particular range of phylotypes is deduced from the sequence data that are available at the time of their design. Due to the exponential growth of sequence databases, some probes may lose their originally envisaged specificity or target group coverage with time. For example, one commonly used probe for Bacteria (EUB338) has been constructed on a set of <1,000 then-available bacterial 16S rRNA gene sequences (6). At the time of this review, the Ribosomal Database Project (42) has collected >170,000 full and partial bacterial sequences. Thus, it is not surprising that the coverage of probe EUB338 is incomplete, and it has been modified accordingly (51). This points to the need to check the target range of existing FISH probes on the latest data set before applying them to unknown samples. A useful database about the current specificity and coverage of many published probes is provided by the Molecular Ecology department of the University of Vienna at http://www.microbial-ecology.de/probebase (162).
However, the replacement of some of the first generation of available FISH probes should be seen as a necessary optimization process in a rapidly progressing field rather than as an irresolvable problem of the FISH approach. Analyses of rRNA gene sequences from aquatic habitats indicate that only a limited number of well-defined phylogenetic clades of microorganisms might actually be common in the pelagic zones of marine and freshwater environments (55, 85, 91, 121, 240, 328). A substantial amount of diversity within several of these clades appears to be covered adequately by the presently available sequence data (108). Such knowledge will eventually provide a reliable base for a new generation of more "habitat-specific" FISH probes that discriminate well-established lineages of microbes in a particular environment (Table 1), e.g., the various marine SAR clades (62, 188, 189, 325) or the freshwater actinobacteria (91, 310).
Another drawback of FISH with fluorescently monolabeled oligonucleotide probes is the low fluorescence intensity of hybridized bacteria from natural water samples (214). Bacteria in oligotrophic water are often small, slow growing, or in stationary phase (187), and their ribosome content is typically low (65). Consequently, there are few rRNA target molecules for FISH staining. The fraction of microbial cells that can be visualized microscopically may thus vary with the physiological state of the studied assemblage. For example, a significantly smaller percentage of bacteria could be stained by FISH in coastal North Sea surface waters during the winter months than during the spring and summer seasons (63). In environments such as offshore marine waters, sometimes only a minor fraction of microbes can be visualized by FISH with fluorescently monolabeled oligonucleotide probes (214). Therefore, it is likely that the abundances of some slowly-growing microorganisms with small cell sizes, e.g., of members of the marine SAR86 clade, are underestimated by the standard FISH approach (210).
During the last decade, efforts have been made to increase the sensitivity of FISH, e.g., with peptide nucleic acid probes (320), brighter fluorochromes (88), image intensified video microscopy (78), preincubation with chloramphenicol (200), hybridization with more than one fluorescently labeled oligonucleotide probe (188), and helper probes (70). Two particularly promising alternative strategies to FISH with fluorescently monolabeled oligonucleotides are polynucleotide probes and enzymatic signal amplification (57, 149, 210). Fluorescently multilabeled rRNA-targeted polyribonucleotide probes yield significantly higher signal intensities than oligonucleotide probes (214). They have been successfully applied to discriminate between bacteria and different groups of archaea in coastal and open ocean environments (40, 136, 214). Limitations of polyribonucleotide probes as a routine tool for the identification of aquatic microbes are the relatively high effort and variability of enzymatic probe synthesis, and the rather low phylogenetic resolution. Only three large taxa have been distinguished by this technique in marine waters (57), although potentially a resolution at the genus level is possible (300).
A working alternative is catalyzed reporter deposition (CARD)-FISH (210, 211, 263), which combines in situ hybridization with horseradish peroxidase labeled oligonucleotide probes and enzymatic signal amplification with fluorescently labeled tyramides (24). This allows the quantitative detection of marine and freshwater pelagic bacteria with low ribosome content that are insufficiently visualized by fluorescently monolabeled probes (210, 263) (Fig. 4).
 View larger version (8K): [in a new window] |
FIG. 4. Comparison of hybridized bacteria from North Sea waters (A) after FISH with fluorescently monolabeled probes and (B) after CARD-FISH. Photographic exposure times were (A) 10 s and (B) 1 s. (Modified from reference 210.)
|
|
TopPreviousNextReferences |
|---|
In order to detect significant net growth or loss of populations, the cell numbers of different microbes in situ have to be determined at a standardized level of precision. This requires manual quantification of the percentage of FISH-stained microbes by epifluorescence microscopy (217). Other potential counting approaches, e.g., flow cytometry (5), so far have failed to provide working alternatives to this tedious evaluation strategy. Eventually, a rapid quantification of bacterial populations by means of automated microscopy (219) might allow us to expand the scale of investigations from a few point measurements to a spatial or temporal resolution that better reflects the true population dynamics of heterotrophic aquatic picoplankton.
A conversion of cell numbers into biomass is required for the reconstruction of carbon fluxes through microbial food webs. Bacterial cell sizes and the relationship between cell size and dry mass can be determined empirically, e.g., by image analyzed microscopy (174, 274) and appropriate conversion factors (159, 197, 233). Some microbial taxa in the plankton form significantly larger cells than others (16), and population changes within such groups may thus contribute disproportionally to changes in total biomass (132, 222). The most extreme example are the filamentous bacteria in the water column of many lakes (130). Such morphotypes rarely form more than a few percent of total cell numbers, but they may temporarily constitute half of total microbial biomass or more (130, 286) (Fig. 5). Therefore, the biomasses of different microbial populations may need to be determined separately in order to assess their respective roles in the carbon flux through aquatic systems (46, 132, 216, 237).
 View larger version (37K): [in a new window] |
FIG. 5. Contribution of filamentous bacteria from the LD2 clade to total bacterial abundances (line and symbols) and biovolume (bars) in the spring plankton of a lake. (Modified from reference 223.)
|
|
TopPreviousNextReferences |
|---|
One important topic of aquatic microbial ecology has been the physiological properties of total microbial assemblages, in particular of the metabolic and respiratory processes that drive biogeochemical transformations (13, 53, 142). In contrast, traditional microbiological research is mainly centered on the physiological properties of bacterial strains in pure culture (109, 127). In between these two levels of analysis there is a conspicuous gap of information. Little is known about the growth-related characteristics of single microbial populations that are realized at environmental conditions in the presence of competitors, predators, viruses, substrate heterogeneity, chemical gradients etc. For example, the genome of the marine planctomycte Rhodopirellula baltica strain 1 contains >100 different sulfatases (90), but it is unclear which ecological advantage is associated with this feature (it might, e.g., reflect in situ growth on a complex mix of substrates such as sulfated polysaccharides). In order to understand the stability or fluctuations of a particular biogeochemical process, it would be important to distinguish if it is mediated by a single, physiologically highly versatile microbial population, or if it is carried out by several bacterial groups that may provide a greater functional redundancy (318).
With the exception of stable isotope analysis (236) or pulse-labeling of nucleic acids (302), physiological information is typically lost by methods that use DNA or rRNA extracts for microbial identification. By contrast, physiological properties can be readily related to a particular genotype at the single cell level. FISH by itself may already provide some information about the physiological state of a population, because the signal intensity of hybridization is proportional to cellular ribosome content. The ribosome content of marine isolates tends to increase with growth rate (138). However, rRNA concentration is a parameter that may sometimes be difficult to interpret. Some marine bacteria may maintain high numbers of ribosomes even during periods of prolonged starvation (68). This is probably essential to rapidly respond to changes in growth conditions in a patchy environment (212). Thus, ribosome content may allow a limited assessment of bacterioplankton "activity" at the community level (52, 80), but it needs to be interpreted with caution if single populations are to be compared. In addition, cell identification by FISH can be combined with a number of other methods that visualize particular physiological properties of individual cells, e.g., substrate uptake, DNA synthesis, respiration (47, 196, 213), and even with stable isotope probing (198).
Microautoradiography. One of the most powerful approaches to study physiological activities in mixed microbial assemblages dates back to the 1960s. The uptake of radiolabeled tracer substrates into individual cells can be visualized by a photographic technique termed microautoradiography (30, 31) (Fig. 6a). In combination with cell identification by FISH, this approach allows us to assess the partitioning of substrate consumption between different microbial populations in mixed assemblages. Microautoradiography-FISH was first used to determine organic and inorganic substrate uptake of ammonia-oxidizing bacteria and of other groups in activated sludge (154), but has since then been adapted for microbes in lakes and in the marine water column (4, 47, 98, 169, 199, 296, 306). Recently, protocols have been developed that integrate the superior CARD-FISH staining with microautoradiography for the analysis of bacterial substrate uptake in environments such as the mesopelagic zone (4, 296). It has even been attempted to add a quantitative aspect to microautoradiography by estimating the amount of incorporated radiolabel from the number of grains that are formed around active cells (46, 168, 170).
 View larger version (70K): [in a new window] |
FIG. 6. (A) Glucose uptake into single cells under anoxic conditions as studied by FISH and microautoradiography. Green objects surrounded by black deposits are Roseobacter sp. cells from coastal North Sea waters that have incorporated radiolabeled substrate. (B) Immunocytochemical detection of DNA de novo synthesis in freshwater bacteria after incubation with bromodeoxyuridine. Green objects: cells affiliated with actinobacteria; yellow-orange objects: bromodeoxyuridine-positive actinobacteria; red objects: other bromodeoxyuridine-positive bacteria. Bar, 10 µm.
|
Fluorescent activity tracers and flow sorting In order to study particular aspects of growth in individual microbial populations there are technically less demanding alternatives to microautoradiography. Pelagic bacteria with an active electron transport system of the respiratory chain reduce tetrazolium salts to water-insoluble crystals (322). Such formazan grains are deposited intracellularly, and they can be detected both microscopically (247) and by flow cytometry (54). In combination with FISH cell respiration can thus be visualized in single microbial populations, as was shown for filamentous bacteria from activated sludge (196). However, the tetrazolium reduction method appears to be a rather insensitive means to distinguish between growing and non-growing (or dead) bacteria in the plankton. Accordingly, bacteria with visible formazan deposits are regarded as the most highly active fraction within a larger set of growing cells (268). While there may be good reasons to identify such highly active bacterial populations, no investigation has combined tetrazolium reduction and FISH to study microbes in the water column of natural aquatic systems.
Bromodeoxyuridine is a halogenated nucleotide analogue of thymidine that is incorporated into newly synthesized DNA of bacteria and eukaryotes (213, 245, 302). It provides a non-radioactive alternative to microautoradiography with tritiated thymidine (207) and it allows us to quantify growth rates at the single-cell level (116). Bromodeoxyuridine incorporation has been combined with CARD-FISH to visualize de novo DNA synthesis in different freshwater and marine bacterial populations (209, 213, 310) (Fig. 6b). This offers a sensitive means to detect changes of growth rates in single microbial populations in situ. During bottle incubations of filtered seawater, a rise in the numbers of bromodeoxyuridine incorporating Alteromonas sp. cells clearly preceded cell multiplication (213). Significant seasonal differences and short-term variability of growth rates were observed in members of the Roseobacter spp. and NOR5 lineages from coastal North Sea picoplankton (209). High bromodeoxyuridine incorporation by actinobacteria in mountain lakes indicated that these bacteria were not passively introduced from soils, but autochthonously growing members of the bacterioplankton community (310).
Flow cytometry may provide an alternative means of detecting activity or substrate uptake in single microbial cells. It allows the physical sorting of particular bacterial populations of interest for further analyses (22, 72). So far, microbial cells from plankton samples have been mainly sorted by phenotypic features, e.g., cell volume or cellular DNA or protein content (72, 150, 267, 326). Sorted bacteria have been analyzed by molecular methods (21, 323) and for radiotracer incorporation (151, 327). In contrast to microautoradiography, the tracer uptake rates of specific cell populations can be readily quantified by this approach.
Direct sorting of microbial cells after FISH staining has first been shown in highly productive wastewater treatment systems (284). Recently this approach has also been adapted for bacterial cells from coastal marine bacterioplankton, taking advantage of the superior signal intensities of CARD-FISH staining (262). Such a combination of cell identification and flow sorting potentially offers the ability to quantitatively investigate substrate uptake of single populations in natural samples (327). Moreover, it might eventually provide a means of obtaining functional genes or larger genome fragments from phylogenetically coherent groups of microbes directly sorted from environmental samples.
For unknown reasons, heterotrophic aquatic microbes form large populations in particular habitats or at particular seasons, and are rare at other locations or time points (35, 63, 136, 144, 218, 280). One challenge of population ecology is to explain the observed distribution patterns of different bacterial taxa from their physiological properties and from their interactions with other organisms. Admittedly this may appear a rather far-fetched goal for a discipline that has just started to understand which microbes are frequent in different aquatic environments. However, the ability of macroecology to understand the role of individual plant and animal species is to a large extent based on an understanding of their distribution patterns and population dynamics at various environmental conditions. So far, only a few studies have investigated abundance changes of particular microbial taxa in the water column in a context of physicochemical parameters or food webs (63, 132, 136, 147, 223, 278).
For the purpose of gaining ecological insight from spatial or temporal distribution patterns, binary information about the presence or absence of a particular bacterial group, as provided by cloning or fingerprinting techniques, is probably insufficient. Such approaches allow us to detect fundamental differences between communities, e.g., between marine and freshwater habitats (180), along rivers (264), or in mesocosm (244, 253), but they can hardly distinguish if a set of environmental variables is more or less favorable for a specific population. A too-coarse division of aquatic microbial assemblages, e.g., into subphyla of Proteobacteria by FISH with the respective probes (171), also suffers from drawbacks, since different populations with potentially contrasting dynamics might be put into ecologically meaningless categories (Fig. 2). Such studies may contribute to our understanding of large biogeographic divisions, e.g., between marine and freshwater habitats (89), and of basic discontinuities in the composition of microbial assemblages, e.g., along estuaries (28). However, both the qualitative community analyses by fingerprinting and investigations on large taxonomic units by FISH should be regarded as intermediate steps towards the quantitative study of ecologically coherent and phylogenetically more tightly-defined populations.
For decades, aquatic microbial ecologists have complemented descriptive studies on the distribution of microbes in various habitats with a range of simple field experiments. These have been referred to as "bottles, bags and buckets" (203), or more respectfully, as micro- to mesocosms, limnocorrals, enclosures, etc. Typically, various volumes of water are taken directly from the environment studied and this water is incubated at more or less in situ conditions after manipulation of, e.g., substrates and nutrients (61, 152, 177, 202), or of particular functional groups of the food web (16, 133, 293, 317). Depending on the container size, the duration of such experiments ranges from days to weeks. Short-term incubations share a basic logic with tracer uptake experiments: a measurable response to a manipulation should allow a deduction about the original state of the assemblage or some of its members. Sometimes dialysis bags with defined pore sizes may provide a more advanced alternative to bottles (80, 172, 278). Such bags allow readier exchange of dissolved substances if exposed directly in the water column. Even so, some features of the original environment are probably irreversibly destroyed, in particular the assumed continuum between the dissolved and the particulate organic matter (12). Larger containers may provide a useful means to artificially induce blooms of specific phytoplankton groups (244) or to manipulate metazooplankton densities (129, 223).
Such investigations are often plagued by the mysterious "bottle effect," a hard-to-define concept that reflects the worry of whether phenomena observed in confined assemblages are nonspecific consequences of the confinement rather than a result of the planned manipulation. Nevertheless, experimental mesocosms are among the few tools available to microbial ecologists that go beyond a purely descriptive analysis of aquatic microbial assemblages. A number of well-defined hypotheses about microbial populations have been successfully addressed by such approaches (16, 61, 132, 223, 244, 278, 293).
Experimental systems such as the Winogradsky column (316) have fundamentally shaped our understanding of environmental microbiology. However, concomitant with the rise of cultivation-independent methods, laboratory investigations on experimental communities have somewhat suffered from a lack of popularity among microbial ecologists. This may have been in parts a consequence of Thomas Brock's harsh words about studies on "mixed cultures of unknown provenance ... at some ill-defined state" (32). It may also be related to the realization that many laboratory investigations have been performed on microorganisms that are readily culturable (112, 212) but that are rare in the water column (62, 312). These drawbacks may no longer apply. For one, an increasing number of aquatic microbes have been isolated during the past years that also form large populations in situ (63, 110, 238) and that thus represent adequate model organisms for laboratory investigations (27, 109). Second, 16S rRNA-based molecular tools now provide new means to precisely follow the population dynamics of different bacteria in mixed experimental assemblages (16, 221). An increasing wealth of genomic information from isolated environmental bacteria may eventually even allow us to link the performances of individual members of such model assemblages to the expression of particular genes in the context of a well-defined experimental setup.
Thus, laboratory investigations on bacterial isolates or model communities add another important dimension to the understanding of microbial population ecology. For example, particular physical structures within aquatic habitats cannot be preserved in experimental approaches in the field. Laboratory systems can artificially produce aggregated organic matter in natural water samples (101) and trap single flocs in a three-dimensional flux field (230). Such designs provide the adequate physical and chemical environment to study microbial activities and successions on organic aggregates and particles (100-102).
Laboratory studies may also provide a better control over parameters that might mask the hypothesized relationship between the studied population and the variable of interest. Continuous-cultivation systems, not necessarily chemostats (126), stabilize the composition of mixed assemblages by enforcing a minimal growth rate and by providing constant temperature, illumination, and input of substrates and nutrients. This allows us to experimentally sustain transient ecological phenomena over prolonged periods of time, e.g., the rise of particular predator populations (112, 221, 232, 234, 250, 279), bacterial-viral interactions (181), or the breakdown of cyanobacterial blooms (304).
Investigations on pure cultures offer the possibility to distinguish between phenotypic and physiological properties of different aquatic bacteria, e.g., in their interactions with algae (29, 99, 146), their chemotactic responses (15, 23), motility patterns (140), grazing sensitivity (27, 114), or cell filamentation (115, 273). This provides the opportunity to test hypotheses about the ecological relevance of such features.
|
TopPreviousNextReferences |
|---|
Our knowledge of aquatic microbial diversity is largely derived from the analysis of 16S rRNA gene sequences directly PCR amplified from environmental DNA (239) and from culture collections predominantly established on rich solid media (107). Both approaches have revealed complementary and often nonoverlapping facets of diversity (but see reference 228). Nevertheless, our perception of the total diversity of aquatic microbes is most probably incomplete. A recent investigation by shotgun cloning of large genome fragments from Sargasso Sea picoplankton concentrates and high-throughput genome analysis has produced >100 novel bacterial 16S rRNA sequence types in a single sample (305).
Evidence has accumulated that there is relatively little overlap between the phylogenetic groups that are present in marine and freshwater environments (89, 180, 241), e.g., specific groups of archaea appear to be entirely limited to the marine water column (59, 76, 173). Lineages of 16 rRNA gene sequence types that occur in both habitat types include the SAR11 clade of Alphaproteobacteria (14, 85, 329), and the ammonia-oxidizing Betaproteobacteria (123, 297). Even within these clades, distinct clusters of marine and freshwater sequence types can be distinguished (329). Some clades of actinobacteria typically found in freshwater habitats also contain sequences from estuaries and marine waters (48, 309), whereas other clades within this phylum are exclusively of either marine or freshwater origin (91, 240, 309, 328). Members of the very diverse Cytophaga/Flexibacter/Flavobacterium group are common in some coastal and offshore marine habitats (45, 63, 144, 280), but they also occur in rivers and lakes (28, 143, 218, 223). Bacteria from this group appear to have radiated across a range of aquatic habitats, including biofilms and sediments (34, 158).
The phylogenetic ties between freshwater and soil microbes are still unclear. Sequence types from both environments have, e.g., been found within the freshwater acIV lineage of the actinobacteria (309). There are furthermore indications that microbial assemblages in some lakes may be similar to those in the influx from the catchment (156).
In the context of this review we distinguish between the multitude of microbial phylogenetic lineages that may occur in various aquatic systems and the few groups of microbes that have been shown to form substantial populations in such environments. Clearly, archival listings of microbial diversity from different habitats are a crucial first step to investigate the role and fate of aquatic microbes, since they provide the essential fundament for subsequent studies about the ecology of particular populations. However, it is equally important to progress from a purely qualitative appreciation of microbial diversity to the quantification of the abundances, biomasses, and activities of different phylogenetic groups. For example, it is presently still unclear if members of the Verrucomicrobia are an important component of freshwater assemblages. Such bacteria are frequently detected in lakes by PCR-based methods (157, 330, 331). From all we know, their densities might be one in a million but potentially also >10% of all cells. The following sections thus specifically discuss investigations that have established the cell concentrations, spatial distributions, temporal successions, or physiological features of specific microbial taxa in the plankton of marine and freshwater systems.
Some groups of marine bacteria had been known for years from their 16S rRNA gene sequences before their abundances in the water column were determined. This is the case for bacteria related to the marine SAR11 (188), SAR86 (210), SAR116 (71), SAR202 (189), and SAR406 (71) clades, whereas for the various lineages of marine actinobacteria (240) such evidence is still lacking. Members of the SAR11 clade are believed to be among the most common prokaryotes in the marine plankton. It has been reported that these bacteria may seasonally represent >50% of total bacterial abundances in surface waters of the northwestern Sargasso Sea and 25% of subeuphotic microbial assemblages (188). Bacteria related to Roseobacter sp. (also referred to as the SAR83 cluster, (241) are another common component of coastal and offshore microbial assemblages, and they may constitute up to 25% of the marine picoplankton (63, 95, 295). The seasonal abundance of Roseobacter spp. in coastal North Sea picoplankton closely followed the development of phytoplankton biomass (Fig. 7) (63, 95, 295). The geographic distribution of one particular subcluster from this lineage appears to be limited to temperate and polar oceans (266). In transects across the German Bight Gammaproteobacteria related to the SAR86 lineage on average formed 7% of total cell numbers (210), and 3 to 6% of all bacterial 16S rRNA genes in Monterey Bay surface waters were affiliated with this group during an upwelling event (295). Genes encoding proteorhodopsin were first described in members of the SAR86 clade (19), but recent findings indicate that such light-driven proton pumps might be a widespread feature of marine bacterioplankton (305).
 View larger version (34K): [in a new window] |
FIG. 7. Seasonal population dynamics of marine Euryarchaeota and of bacteria from the Roseobacter lineage in the German Bight of the North Sea. Bars: phytoplankton biomass. (Combined from references 63 and 214.)
|
The composition of microbial communities in more extreme habitats might sometimes be very simple. Hydrothermal circulation activities in deep sea environments produce buoyant plumes with substantially elevated levels of reduced chemicals. The bacterial assemblages within such a plume inside the caldera of the Suiyo Seamount volcano consisted almost entirely of two distinct phylogenetic populations that were related to sulfur-oxidizing symbionts of hydrothermal vent fauna (291). The caldera might thus represent a giant natural continuous-cultivation system for these two groups.
Sometimes, it may be necessary to define microbial populations at the level of single strains, e.g., for the study of pathogenic Vibrio spp. in marine waters. In coastal environments Vibrio cholerae can be found both attached on particles and free-living in the water column (44), and pronounced seasonal and horizontal differences in population sizes have been reported (83, 117). In microcosms spiked with V. cholerae, rapid growth of these bacteria was observed after addition of organic carbon (190). It is thus conceivable that anthropogenic eutrophication might indirectly favor the growth and dispersal of pathogenic Vibrio strains.
The two major pelagic lineages of Crenarchaeota and Euryarchaeota are among the most well studied phylogenetic group of uncultured microbes in marine picoplankton. Both oligo- and polynucleotide probes have been developed for the direct visualization of such microbes in water samples (57, 78, 192, 296). Recently, a protocol for CARD-FISH staining of Archaea in samples from the deep sea has been described (296). Originally it was believed that creanarchaeota only form large populations in the meso- and bathypelagic layers below the euphotic zone (59, 76). In a seasonal study in the North Pacific subtropical gyre, the mean annual abundances of Crenarchaeota below 200 m water depth ranged between 20 and 40% of total picoplankton cells, which corresponded to 3 x 103 to 1 x 105 cells ml–1 (136). Comparable abundances of this archaeal group were also reported from the deeper waters of the Antarctic circumpolar deep water (40). However, the same study also detected large crenarchaeal populations by FISH in the surface waters of the Southern Ocean during the winter months. Such contrasting vertical distribution patterns are currently difficult to interpret.
The metabolic capacities of planktonic crenarchaea are unclear, but there are indications that members of this lineage might be auto- or mixotrophic (120, 321). The planktonic marine Euryarchaeota on the other hand appear to be a common element of coastal assemblages and surface waters. Members of this lineage seasonally formed >30% of all cells in the surface picoplankton of the North Sea (214) (Fig. 7). Seasonal blooms of Euryarchaeota were also observed during a long-term study in surface waters of the upper Santa Barbara Channel (191).
Seasonal dynamics of different freshwater bacterioplankton populations have first been reported from an ultraoligotrophic mountain lake (218). So far, bacteria from two of the four typical freshwater lineages of Betaproteobacteria as defined by Glöckner et al. (91) have been detected in high abundance in the environment. Bacteria affiliated with the beta I clade (also termed the "Rhodoferax sp. BAL47 lineage") (328) formed populations of >10% in the summer plankton of a eutrophic reservoir (278). A second lineage of Betaproteobacteria related to Polynucleobacter necessarius (beta II) seasonally constituted up to 50% of all pelagic microbes in the aerobic waters of a meromictic humic lake (35). Filamentous bacteria from the LD2 subclade (328) that is closely related to Haliscomenobacter hydrossis (255) (Cytophaga/Flexibacter/Flavobacterium) transiently formed >40% of total bacterial biomass in a mesotrophic lake (223). Actinobacteria from the uncultured acI clade (91, 309, 328) are another ubiquitous group of freshwater prokaryotes (91, 309, 328) that seasonally occur in high densities in habitats of very different limnological characteristics, e.g., humic (35) or high mountain lakes (91, 263, 310).
Microbial assemblages on suspended organic aggregates differ from those of the water column (56). So far the particle-attached communities in marine habitats have only been investigated qualitatively by fingerprinting and comparative sequence analysis (48, 56). More information is available about limnic and riverine aggregates. Microbial populations on aggregates in fresh waters change with the ageing of such particles (101) or during their transport from rivers into estuaries (265). Three populations related to the genera Duganella, Hydrogenophaga, and Acidovorax formed almost half of the Betaproteobacteria on organic aggregates obtained from Lake Constance at a depth of 50 m (261). These bacteria effectively colonized artificially produced microaggregates within 24 h. Interestingly, members of the three genera were rarely detected in the planktonic microbial assemblage, and they were not affiliated with the presently defined clades of typical freshwater Betaproteobacteria (91, 329). Instead, these bacteria are known from highly eutrophic environments such as activated sludge (249, 258).
In contrast to the lake snow assemblages, riverine organic particles have been described to mainly harbor Betaproteobacteria related to the drinking water biofilm bacterium Aquabacterium commune. These bacteria formed up to 30% of all cells on lotic organic aggregates in the river Elbe (135). The composition of the microbial assemblages on such aggregates showed pronounced seasonal changes, and bacteria related to the Planctomycetales were absent in winter.
|
TopReferences |
|---|
The shortness of the following section does not reflect the importance we assign to the topic. Our current understanding of the relationship between the growth of individual bacterial populations and the availability of particular substrates or nutrients is still rudimentary. For example, it is established that pelagic bacteria and archaea may incorporate amino acids (201), but with a few exceptions (see below) it is unclear if different genotypes show preferences for individual components of such mixes. Studies on the interactions between bacteria and specific algae indirectly suggest that substrate quality may play a role in the competition between microbial populations. Pure cultures of various algal species may harbor distinct microbial assemblages (252). Different microbial communities could be established from identical inocula in seawater micrososms if blooms of specific algal groups were induced (225). Cottrell and Kirchman reported (47) that various cooccurring microbial populations in pelagic habitats may be specialized on the degradation and uptake of particular substrate classes. Using microautoradiography, these authors showed that different bacterial genotypes also differed in their uptake of 3H from radiolabeled chitin, proteins, amino acids, and N-acetylglucosamine. A preference for simple monomers over proteins in bacteria from the SAR11 clade has recently been reported by the same laboratory (168).
A set of recent investigations on the transfer and processing of phytoplankton-derived organic sulfur compounds has provided a first model for future studies of the relationship between particular bacterial populations and a specific biogeochemical process. Dissolved dimethylsulfonopropionate (dDMSP) is an algal osmolyte that is released during viral lysis or sloppy zooplankton feeding (50, 167). dDMSP is a growth substrate for heterotrophic bacteria (153), and it represents an important precursor of protein-sulfur in marine bacterioplankton (139). An increasing body of evidence is pointing to a few groups of marine bacteria as key elements in dDMSP biogeochemistry. Laboratory studies on isolates of the diverse Roseobacter clade have indicated a widespread ability of members of this lineage to degrade dDMSP and to incorporate the sulfur derived from this organic compound into cellular protein (93). In addition, large populations of bacteria affiliated with the Roseobacter clade were observed during dDMSP-producing algal blooms in the North Atlantic. In two independent studies the horizontal or vertical distribution of microbes from this group was positively correlated to dDMSP concentrations or fluxes (96, 323). Direct evidence of DMSP uptake was recently obtained by microautoradiography (170, 306). This approach also revealed that DMSP uptake may be a feature of different bacterial populations, and bacteria from the SAR11 clade rather than Roseobacter mediated the bulk of DMSP turnover in an offshore planktonic assemblage (169).
Selective preference for particular substrates may not be the only factor affecting the competition between aquatic microbes. Specialization on a single resource might in fact be disadvantageous in an energy-deficient environment (60), and the majority of bacteria and archaea in pelagic marine environments are capable of incorporating mixes of radiolabeled amino acids (137).
A considerable proportion of the substrates and of bacterial productivity in freshwater and coastal habitats are distributed in organic particles and microscale patches (12). In contrast, the bathypelagic zone of the open oceans represents a rather desert-like environment poor in particulate organic matter. Individual microbial species or phylogenetic lineages within the bacterioplankton thus likely differ in their ability to succeed in habitats with steeper or flatter substrate gradients. An "opportunistic" growth strategy might be widespread among those bacterial groups that successfully colonize organic aggregates or other nutrient-rich microniches. A marine Pseudoalteromonas sp. strain exhibited a significantly shorter growth delay than an Oceanospirillum sp. under "feast-and-famine" batch culture conditions, but the former strain experienced a growth disadvantage if substrate changes were gradual (212). The importance of such bacteria may vary between habitats, and different species of "opportunistically" growing microbes are found in marine and freshwater environments (224).
Representatives of "opportunistic" genera of Gammaproteobacteria that are readily culturable on solid media were generally rare in the bacterioplankton of shallow North Sea waters (62). In contrast, colony-forming bacteria represented a prominent fraction of the microbial assemblages in brackish waters of the coastal Baltic Sea (227, 228). Readily enrichable bacteria were also found in high concentrations in the plankton of a lake with high irregular input of organic carbon from the catchment (35). The identification and quantification of such "r-strategists" within heterotrophic picoplankton assemblages might thus eventually allow deductions about short-term fluctuations in the availability of organic matter in aquatic habitats, e.g., for a biological monitoring of pollution effects. In contrast, many if not most of the typical free-living marine and freshwater bacteria appear to lack the ability to form colonies on solid media (282), and their growth is negatively affected by enhanced substrate levels (289).
It is likely that many bacterial species in the water column combine elements of the oligotrophic and the "opportunistic" growth strategies in their life cycles, e.g., by alternating between free-living and attached growth forms (141). Isolates from freshwater plankton that were initially oligocarbophilic could gradually be adapted to richer conditions (114). Bacteria from the marine NOR5 lineage, which are apparently free-living in the water column, also formed colonies on low-nutrient agar plates, albeit significantly later than other "opportunistic" strains (63). A dual-niche existence may also explain why facultatively anaerobic bacteria appear to be such a common component of coastal bacterioplankton assemblages (4, 243). In shallow waters of the North Sea >80% of free-living Roseobacter spp. cells were able to incorporate glucose at both oxic and anoxic conditions (4). Since the average depth of the German Bight is only 20 m, bacteria from the Roseobacter spp. lineage that colonize aggregated senescent algae (97) might experience temporary anoxia while settled on the sediment surface. These bacteria could be reintroduced into the water column by the periodic resuspension of particulate organic matter (179).
So far there is little evidence that a lack of resources is a major cause of bacterioplankton mortality. Cultured strains that are thought to be representative for the pelagic environment are often oligocarbophilic (238, 259), or they survive extended periods of starvation (68). Predation and viral lysis are believed to be the key factors that counterbalance microbial growth in the water column (13, 77, 271). Growth inhibition or cell damage induced by UV radiation may be another important ecological factor in some aquatic habitats (10, 119, 285). Nevskia ramosa, a species inhabiting the neuston layer of fresh waters, exhibited elevated resistance to UV (290). Some alpine lakes exposed to intense levels of UV-B radiation (148) feature conspicuously high abundances of actinobacteria, a phylogenetic lineage of mainly gram-positive bacteria with a high genomic G+C content (91, 263, 310). Gram-positive bacterial isolates are often less affected by UV radiation than gram-negative bacteria (8), and a high genomic G+C content has been suggested to mediate higher resistance to radiation damage (166).
Viral lysis. The current concepts of specific viral-bacterial interaction in aquatic systems are largely shaped by theoretical models (298), and presently there are almost no investigations about the influence of viral lysis on the coexistence of individual microbial populations in the plankton (260, 317). Viral influence on the growth of different bacteria might act both directly by "killing the winner" (298), but also indirectly via the release of dissolved organic matter and nutrients from lysed pro- and eukaryotic cells (92, 182). Rapid development of resistance to viral infection has been observed in some bacterial strains during continuous-culture experiments (181), but it is unclear if such a process will also occur in natural aquatic assemblages. Recently, changes of the species richness of marine archaea have been reported as a likely consequence of experimentally manipulating viral densities (317). For a detailed discussion of the influence of viruses on aquatic microbial assemblages, readers should refer to specific reviews on the subject (311, 319).
Selective predation. Hetero- and mixotrophic protists, in particular nanoflagellates and ciliates, are the main consumers of picoplankton in the marine and freshwater pelagic zone. Their role in controlling the abundance, biomass, and productivity of microbial assemblages has been amply documented (269, 298). Many of these predators are omnivorous, i.e., they can feed on a large range of bacterial species. However, they are not unselective feeders. In the following we will focus on predation selectivity that is related to prey cell morphology, in order to illustrate the influence of a phenotypic feature on the success of different genotypic microbial populations. Other aspects of microbial predator-prey interactions are discussed elsewhere (25, 82, 131, 215, 277).
Due to their specific particle uptake mechanisms and handling procedures, protistan predators cannot feed on all bacterial shapes or cell size classes with equal efficiency (26, 178). As a consequence, microbial cells within a length range of 1 to 3 µm are preferably ingested by heterotrophic flagellates and ciliates, whereas smaller or larger cells profit from reduced loss rates (130, 186, 276). Even small differences in cell sizes between strains of freshwater ultramicrobacteria may have large effects on their survival rates (27). Microbial assemblages that are exposed to high grazing pressure thus typically harbor high abundances of extremely small cells (222). In contrast, large filamentous bacterial morphotypes and bacteria that form microcolonies are substantially more protected from protistan grazing (115, 273). Filamentous morphotypes accumulate during periods of high predation in fresh waters (134), but they are rarely observed in marine systems. Some strains of freshwater Betaproteobacteria produce microcolonies within a sponge-like matrix of extrapolymerous substances that renders them resistant to flagellate predation (113).
Size-selective predation induces shifts in the genotypic composition of mixed assemblages by imposing different mortality rates on bacterial species with different mean cell sizes (Fig. 8) (221). First evidence for such community changes originated from continuous-cultivation experiments on microbial assemblages that formed stable associations with freshwater algae (221, 234). Selective grazing mortality is moreover believed to set an upper limit to the standing stocks of some microbial populations in the plankton. For example, bacteria from a cosmopolitan lineage of freshwater Betaproteobacteria (beta I) constituted approximately 10% of the summer assemblage in a eutrophic drinking-water reservoir (278). After removal of protists and in situ incubations in dialysis bags, members of the beta I clade increased to almost 30% of total cells within 24 h. It is thus likely that these bacteria contributed disproportionally to the flux of organic carbon from the picoplankton to the higher trophic levels. Some genera of "opportunistically" growing Gammaproteobacteria (Alteromonas, Vibrio, and Pseudoalteromonas) are probably rare in coastal surface picoplankton because they are almost completely suppressed by size-selective grazing (16). This mechanism might also play a role in eliminating pathogenic Vibrio spp. from the water column (165).
 View larger version (32K): [in a new window] |
FIG. 8. Effect of size-selective protistan grazing on bacterioplankton biomass size distribution. Several microbial populations with different mean cell sizes are highlighted. (a) Low or no grazing: unimodal cell size distribution. Most biomass is distributed in populations close to the mean size, whereas very small or large populations are rare. (b) Heavy grazing: populations within the edible cell size range are eliminated unless they can escape predation by a change in their mean cell sizes. Under these conditions the growth of very small or very large phenotypes is favored. (Reproduced with permission from Nature Reviews Microbiology [215] copyright Macmillan Magazines Ltd.)
|
The interplay between resource availability and mortality in determining the population sizes of different bacterial taxa is poorly understood. Changes in bacterial community composition might be related to changes in the ratio of bacterial mortality rate to growth rate (277). This would imply that communities with apparently stable taxonomic composition may be encountered at very different levels of total microbial productivity or mortality and that profound community changes are induced by shifts from top down to bottom up or vice versa (235). One type of natural model system to study such interactions might be freshwater reservoirs that feature pronounced longitudinal substrate and nutrient gradients between the river influx and the dam areas. Experiments that combine the transplant of water from different positions of the gradient with food web manipulations (80) might eventually shed more light on the complex control of different microbial populations by top-down and bottom-up forces.
The future of environmental microbiology will be shaped by the current advances in genome sequence analysis of cultured and uncultured microbes. This appears to create a favorable conceptual environment for a population-based perspective in aquatic microbial ecology. Already, the analysis of genomic information has strengthened the interest in the cultivation of abundant heterotrophic microbes from the water column (39, 63, 110, 204, 238). Such activities will probably draw increased attention to the unique physiological features of different genotypes and thus to the potential ecological differences between cooccurring microbial populations in mixed assemblages.
Some genomic information will moreover doubtlessly be of direct interest for the study of aquatic microbial populations in situ. Emerging techniques such as a combined FISH staining of mRNA and rRNA (208) will allow us to detect the expression of metabolic genes within bacterioplankton populations defined at the level of single ecotypes (41). Ideally, such gene-specific probing could be applied in combination with other ecologically relevant features of bacterial cells, e.g., morphology (223), proliferation (213), substrate uptake and turnover (324), motility (104), exoenzyme activity (195), or polysaccharide sheath formation (288). Moreover, a more complete knowledge of microbial genomes will allow us to detect far more subtle phylogenetic differences between bacteria than what can presently be derived from the analysis of 16S rRNA genes (315). Questions addressing the biogeography (43, 315) and physiological or evolutionary adaptation of microbial ecotypes (27, 109, 128) might therefore form another area of future research in environmental microbiology.
However, the ties between environmental genomics and microbial population ecology should not be overstated. The population sizes of different microbes in the environment cannot be predicted from their genomic constitutions only, and the success of different bacterial taxa is codetermined by their interactions with competitors and predators in the context of the physicochemical environment. Moreover, while the existence of microbial biogeography arguably is debatable (66), this controversy cannot be a central concern of population ecology. Even if in principle "everything is everywhere" (18), it is known that different groups of heterotrophic aquatic microbes exhibit distinct temporal, spatial, and geographic distribution optima. Thus, a major challenge of the coming decade will be to understand the underlying reasons for the heterogenous distribution patterns of different microbial populations in marine and freshwater plankton.
We thank our students and colleagues for all the fruitful discussions and controversies that helped to shape our ideas and David Kirchman and two anonymous reviewers for their helpful comments on the manuscript.
We acknowledge the institutions that have supported our research on the topics discussed here, in particular the European Union (EVK3-2001-00194 BASICS), the German Ministry of Education and Research (BMBF01 LC0021/TP4), the German Research Foundation (PE-918/2-1), and the Max Planck Society.
* Corresponding author. Present address: Limnological Station, Institute of Plant Biology, Seestrasse 187, CH-8802 Kilchberg, Switzerland. Phone: 41-1-716-1210. E-mail: pernthaler{at}limnol.unizh.ch.
|
Top |
|---|
- Acinas, S. G., V. Klepac-Ceraj, D. E. Hunt, C. Pharino, I. Ceraj, D. L. Distel, and M. F. Polz. 2004. Fine-scale phylogenetic architecture of a complex bacterial community. Nature 430:551-554.[CrossRef][Medline]
- Alfreider, A., J. Pernthaler, R. Amann, B. Sattler, F.-O. Glöckner, A. Wille, and R. Psenner. 1996. Community analysis of the bacterial assemblages in the winter cover and pelagic layers of a high mountain lake by in situ hybridization. Appl. Environ. Microbiol. 62:2138-2144.[Abstract]
- Alldredge, A., and M. Silver. 1988. Characteristics, dynamics and significance of marine snow. Prog. Oceanogr. 20:41-82.[CrossRef]
- Alonso, C., and J. Pernthaler. 2005. Incorporation of glucose at anoxic conditions by bacterioplankton from coastal North Sea surface waters. Appl. Environ. Microbiol. 71:1709-1716.
[Abstract/Free Full Text] - Amann, R. I., B. J. Binder, R. J. Olson, S. W. Chisholm, R. Devereux, and D. A. Stahl. 1990. Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Appl. Environ. Microbiol. 56:1919-1925.
[Abstract/Free Full Text] - Amann, R. I., L. Krumholz, and D. A. Stahl. 1990. Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J. Bacteriol. 172:762-770.
[Abstract/Free Full Text] - Amann, R. I., W. Ludwig, and K. H. Schleifer. 1995. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59:143-169.[Abstract]
- Arrage, A. A., T. J. Phelps, R. E. Benoit, and D. C. White. 1993. Survival of subsurface microorganisms exposed to UV radiation and hydrogen peroxide. Appl. Environ. Microbiol. 59:3545-3550.
[Abstract/Free Full Text] - Arrieta, J. M., and G. I. Herndl. 2002. Changes in bacterial beta-glucosidase diversity during a coastal phytoplankton bloom. Limnol. Oceanogr. 47:594-599.
- Arrieta, J. M., M. G. Weinbauer, and G. J. Herndl. 2000. Interspecific variability in sensitivity to UV radiation and subsequent recovery in selected isolates of marine bacteria. Appl. Environ. Microbiol. 66:1468-1473.
[Abstract/Free Full Text] - Azam, F. 2001. Introduction, history, and overview: the ‘methods’ to our madness. Methods Microbiol. 30:1-12.[CrossRef]
- Azam, F. 1998. Microbial control of oceanic carbon flux: the plot thickens. Science 280:694-696.
[Free Full Text] - Azam, F., T. Fenchel, J. G. Field, J. S. Gray, L. A. Meyer-Reil, and F. Thingstad. 1983. The ecological role of water-column microbes in the sea. Mar. Ecol. Prog. Ser. 10:257-263.
- Bahr, M., J. E. Hobbie, and M. L. Sogin. 1996. Bacterial diversity in an arctic lake: a freshwater SAR11 cluster. Aquat. Microb. Ecol. 11:271-277.
- Barbara, G. M., and J. G. Mitchell. 2003. Marine bacterial organisation around point-like sources of amino acids. FEMS Microbiol. Ecol. 43:99-109.[CrossRef]
- Beardsley, C., J. Pernthaler, W. Wosniok, and R. Amann. 2003. Are readily cultured bacteria in coastal North Sea waters suppressed by selective grazing mortality? Appl. Environ. Microbiol. 69:2624-2630.
[Abstract/Free Full Text] - Becker, S., M. Fahrbach, P. Boger, and A. Ernst. 2002. Quantitative tracing, by Taq nuclease assays, of a Synechococcus ecotype in a highly diversified natural population. Appl. Environ. Microbiol. 68:4486-4494.
[Abstract/Free Full Text] - Beijerinck, M. W. 1913. De infusies en de ontdekking der backteriën. Müller, Amsterdam, The Netherlands.
- Beja, O., L. Aravind, E. V. Koonin, M. T. Suzuki, A. Hadd, L. P. Nguyen, S. Jovanovich, C. M. Gates, R. A. Feldman, J. L. Spudich, E. N. Spudich, and E. F. DeLong. 2000. Bacterial rhodopsin: evidence for a new type of phototrophy in the sea. Science 289:1902-1906.
[Abstract/Free Full Text] - Beja, O., E. N. Spudich, J. L. Spudich, M. Leclerc, and E. F. DeLong. 2001. Proteorhodopsin phototrophy in the ocean. Nature 411:786-789.[CrossRef][Medline]
- Bernard, L., C. Courties, C. Duperray, H. Schafer, G. Muyzer, and P. Lebaron. 2001. A new approach to determine the genetic diversity of viable and active bacteria in aquatic ecosystems. Cytometry 43:314-321.[CrossRef][Medline]
- Bernard, L., C. Courties, P. Servais, M. Troussellier, M. Petit, and P. Lebaron. 2000. Relationships among bacterial cell size, productivity, and genetic diversity in aquatic environments using cell sorting and flow cytometry. Microb. Ecol. 40:148-158.[Medline]
- Blackburn, N., T. Fenchel, and J. Mitchell. 1998. Microscale nutrient patches in planktonic habitats shown by chemotactic bacteria. Science 282:2254-2256.
[Abstract/Free Full Text] - Bobrow, M. N., T. D. Harris, K. J. Shaughnessy, and G. J. Litt. 1989. Catalyzed reporter deposition, a novel method of signal amplification—application to immunoassays. J. Immunol. Methods 125:279-285.[CrossRef][Medline]
- Boenigk, J., and H. Arndt. 2002. Bacterivory by heterotrophic flagellates: community structure and feeding strategies. Antonie van Leeuwenhoek 81:465-480.[CrossRef][Medline]
- Boenigk, J., and H. Arndt. 2000. Particle handling during interception feeding by four species of heterotrophic nanoflagellates. J. Eukaryot. Microbiol. 47:350-358.[CrossRef][Medline]
- Boenigk, J., P. Stadler, A. Wiedlroither, and M. W. Hahn. 2004. Strain-specific differences in the grazing sensitivities of closely related ultramicrobacteria affiliated with the Polynucleobacter cluster. Appl. Environ. Microbiol. 70:5787-5793.
[Abstract/Free Full Text] - Bouvier, T. C., and P. A. del Giorgio. 2002. Compositional changes in free-living bacterial communities along a salinity gradient in two temperate estuaries. Limnol. Oceanogr. 47:453-470.
- Bratbak, G., and T. F. Thingstad. 1985. Phytoplankton bacteria interactions—an apparent paradox. Analysis of a model system with both competition and commensalism. Mar. Ecol. Prog. Ser. 25:23-30.
- Brock, M. L., and T. D. Brock. 1968. The application of micro-autoradiographic techniques to ecological studies. Mitt. Int. Verein. Limnol. 15:1-29.
- Brock, T. D. 1967. Bacterial growth rate in the sea - direct analysis by thymidine autoradiography. Science 155:81-83.
[Abstract/Free Full Text] - Brock, T. D. 1987. The study of microorganisms in situ: progress and problems, p. 1-17. In M. Fletcher, T. R. G. Gray, and J. G. Jones (ed.), Ecology of microbial communities. Cambridge University Press, Cambridge, England.
- Brosius, J., T. J. Dull, D. D. Sleeter, and H. F. Noller. 1981. Gene organization and primary structure of a ribosomal RNA operon from Escherichia coli. J. Mol. Biol. 148:107-127.[CrossRef][Medline]
- Brümmer, I. H. M., W. Fehr, and I. Wagner-Döbler. 2000. Biofilm community structure in polluted rivers: abundance of dominant pylogenetic groups over a complete annual cycle. Appl. Environ. Microbiol. 66:3078-3082.
[Abstract/Free Full Text] - Burkert, U., F. Warnecke, B. H.-D., E. Zwirnmann, and J. Pernthaler. 2003. Members of a readily enriched beta-proteobacterial clade are common in the surface waters of a humic lake. Appl. Environ. Microbiol. 69:6550-6559.
[Abstract/Free Full Text] - Casamayor, E. O., R. Massana, S. Benlloch, L. Ovreas, B. Diez, V. J. Goddard, J. M. Gasol, I. Joint, F. Rodriguez-Valera, and C. Pedros-Alio. 2002. Changes in archaeal, bacterial and eukaryal assemblages along a salinity gradient by comparison of genetic fingerprinting methods in a multipond solar saltern. Environ. Microbiol. 4:338-348.[CrossRef][Medline]
- Chin, K. J., T. Lukow, and R. Conrad. 1999. Effect of temperature on structure and function of the methanogenic archaeal community in an anoxic rice field soil. Appl. Environ. Microbiol. 65:2341-2349.
[Abstract/Free Full Text] - Chisholm, S. W., R. J. Olson, E. R. Zettler, R. Goericke, J. B. Waterbury, and N. A. Welschmeyer. 1988. A novel free-living prochlorophyte abundant in the oceanic euphotic zone. Nature 334:340-343.[CrossRef]
- Cho, J. C., and S. J. Giovannoni. 2004. Cultivation and growth characteristics of a diverse group of oligotrophic marine gammaproteobacteria. Appl. Environ. Microbiol. 70:432-440.
[Abstract/Free Full Text] - Church, M. J., E. F. DeLong, H. W. Ducklow, M. B. Karner, C. M. Preston, and D. M. Karl. 2003. Abundance and distribution of planktonic Archaea and Bacteria in the waters west of the Antarctic Peninsula. Limnol. Oceanogr. 48:1893-1902.
- Cohan, F. M. 2002. What are bacterial species? Annu. Rev. Microbiol. 56:457-487.[CrossRef][Medline]
- Cole, J. R., B. Chai, T. L. Marsh, R. J. Farris, Q. Wang, S. A. Kulam, S. Chandra, D. M. McGarrell, T. M. Schmidt, G. M. Garrity, and J. M. Tiedje. 2003. The Ribosomal Database Project (RDP-II): previewing a new autoaligner that allows regular updates and the new prokaryotic taxonomy. Nucleic Acids Res. 31:442-443.
[Abstract/Free Full Text] - Colwell, R. R., and A. Huq. Global microbial ecology: biogeography and diversity of vibrios as a model. J. Appl. Microbiol. 85:134S-137S, 1999.
- Colwell, R. R., R. J. Seidler, J. Kaper, S. W. Joseph, S. Garges, H. Lockman, D. Maneval, H. Bradford, N. Roberts, E. Remmers, I. Huq, and A. Huq. 1981. Occurrence of Vibrio cholerae serotype-O1 in Maryland and Louisiana estuaries. Appl. Environ. Microbiol. 41:555-558.
[Abstract/Free Full Text] - Cottrell, M. T., and D. L. Kirchman. 2000. Community composition of marine bacterioplankton determined by 16S rRNA gene clone libraries and fluorescence in situ hybridization. Appl. Environ. Microbiol. 66:5116-5122.
[Abstract/Free Full Text] - Cottrell, M. T., and D. L. Kirchman. 2003. Contribution of major bacterial groups to bacterial biomass production (thymidine and leucine incorporation) in the Delaware estuary. Limnol. Oceanogr. 48:168-178.
- Cottrell, M. T., and D. L. Kirchman. 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.
[Abstract/Free Full Text] - Crump, B. C., E. V. Armbrust, and J. A. Baross. 1999. Phylogenetic analysis of particle-attached and free-living bacterial communities in the Columbia River, its estuary, and the adjacent coastal ocean. Appl. Environ. Microbiol. 65:3192-3204.
[Abstract/Free Full Text] - Crump, B. C., G. W. Kling, M. Bahr, and J. E. Hobbie. 2003. Bacterioplankton community shifts in an arctic lake correlate with seasonal changes in organic matter source. Appl. Environ. Microbiol. 69:2253-2268.
[Abstract/Free Full Text] - Dacey, J. W. H., and S. G. Wakeham. 1986. Oceanic dimethylsulfide-production during zooplankton grazing on phytoplankton. Science 233:1314-1316.
[Abstract/Free Full Text] - Daims, H., A. Bruhl, R. Amann, K. H. Schleifer, and M. Wagner. 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.[Medline]
- del Giorgio, P. A., and T. C. Bouvier. 2002. Linking the physiologic and phylogenetic successions in free- living bacterial communities along an estuarine salinity gradient. Limnol. Oceanogr. 47:471-486.
- del Giorgio, P. A., J. J. Cole, and A. Cimbleris. 1997. Respiration rates in bacteria exceed phytoplankton production in unproductive aquatic systems. Nature 385:148-151.[CrossRef]
- del Giorgio, P. A., Y. T. Prairie, and D. F. Bird. 1997. Coupling between rates of bacterial production and the abundance of metabolically active bacteria in lakes, enumerated using CTC reduction and flow cytometry. Microb. Ecol. 34:144-154.[CrossRef][Medline]
- DeLong, E. F. 1992. Archaea in coastal marine environments. Proc. Natl. Acad. Sci. USA 89:5685-5689.
[Abstract/Free Full Text] - DeLong, E. F., D. G. Franks, and A. L. Alldredge. 1993. Phylogenetic diversity of aggregate-attached vs. free-living marine bacterial assemblages. Limnol. Oceanogr. 38:924-934.
- DeLong, E. F., L. T. Taylor, T. L. Marsh, and C. M. Preston. 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.
[Abstract/Free Full Text] - DeLong, E. F., G. S. Wickham, and N. R. Pace. 1989. Phylogenetic strains: ribosomal RNA-based probes for the identification of single cells. Science 243:1360-1363.
[Abstract/Free Full Text] - DeLong, E. F., K. Y. Wu, B. B. Prezelin, and R. V. M. Jovine. 1994. High abundance of Archaea in Antarctic marine picoplankton. Nature 371:695-697.[CrossRef][Medline]
- Egli, T. 1993. The ecological and physiological significance of the growth of heterotrophic microorganisms with mixes of substrates, p. 305-385. In J. G. Jones (ed.), Advances in microbial ecology, vol. 13. Plenum Press, New York, N.Y.
- Eilers, H., J. Pernthaler, and R. Amann. 2000. Succession of pelagic marine bacteria during enrichment: A close look on cultivation-induced shifts. Appl. Environ. Microbiol. 66:4634-4640.
[Abstract/Free Full Text] - Eilers, H., J. Pernthaler, F. O. Glöckner, and R. Amann. 2000. Culturability and in situ abundance of pelagic bacteria from the North Sea. Appl. Environ. Microbiol. 66:3044-3051.
[Abstract/Free Full Text] - Eilers, H., J. Pernthaler, J. Peplies, F. O. Glöckner, G. Gerdts, and R. Amann. 2001. Isolation of novel pelagic bacteria from the German Bight and their seasonal contribution to surface picoplankton. Appl. Environ. Microbiol. 67:5134-5142.
[Abstract/Free Full Text] - Elena, S. F., and R. E. Lenski. 2003. Evolution experiments with microorganisms: The dynamics and genetic bases of adaptation. Nat Rev. Gen. 4:457-469.[CrossRef][Medline]
- Fegatella, F., J. Lim, S. Kjelleberg, and R. Cavicchioli. 1998. Implications of rRNA operon copy number and ribosome content in the marine oligotrophic ultramicrobacterium Sphingomonas sp. strain Rb2256. Appl. Environ. Microbiol. 64:4433-4438.
[Abstract/Free Full Text] - Fenchel, T. 2003. Biogeography for bacteria. Science 301:925-926.
[Abstract/Free Full Text] - Ferris, M. J., and B. Palenik. 1998. Niche adaptation in ocean cyanobacteria. Nature 396:226-228.[CrossRef]
- Flärdh, K., P. S. Cohen, and S. Kjelleberg. 1992. Ribosomes exist in large excess over the apparent demand for protein synthesis during carbon starvation in marine Vibrio sp. strain CCUG 15956. J. Bacteriol. 174:6780-6788.
[Abstract/Free Full Text] - Frostegard, A., S. Courtois, V. Ramisse, S. Clerc, D. Bernillon, F. Le Gall, P. Jeannin, X. Nesme, and P. Simonet. 1999. Quantification of bias related to the extraction of DNA directly from soils. Appl. Environ. Microbiol. 65:5409-5420.
[Abstract/Free Full Text] - Fuchs, B. M., F. O. Glöckner, J. Wulf, and R. Amann. 2000. Unlabeled helper oligonucleotides increase the in situ accessibility to 16S rRNA of fluorescently labeled oligonucleotide probes. Appl. Environ. Microbiol. 66:3603-3607.
[Abstract/Free Full Text] - Fuchs, B. M., D. Woebken, M. V. Zubkov, P. Burkill, and R. Amann. 2005. Molecular identification of picoplankton populations in contrasting waters of the Arabian Sea. Aquat. Microb. Ecol. 39:145-157.
- Fuchs, B. M., M. V. Zubkov, K. Sahm, P. H. Burkill, and R. Amann. 2000. Changes in community composition during dilution cultures of marine bacterioplankton as assessed by flow cytometric and molecular biological techniques. Environ. Microbiol. 2:191-201.[CrossRef][Medline]
- Fuhrman, J. A. 2002. Community structure and function in prokaryotic marine plankton. Antonie van Leeuwenhoek 81:521-527.[CrossRef][Medline]
- Fuhrman, J. A., and F. Azam. 1980. Bacterioplankton secondary production estimates for coastal waters of British Columbia, Antarctica, and California. Appl. Environ. Microbiol. 39:1085-1095.
[Abstract/Free Full Text] - Fuhrman, J. A., D. E. Comeau, A. Hagström, and A. M. Chan. 1988. Extraktion from natural planctonic microorganisms of DNA suitable for molecular biological studies. Appl. Environ. Microbiol. 54:1426-1429.
[Abstract/Free Full Text] - Fuhrman, J. A., K. McCallum, and A. A. Davis. 1992. Novel major archaebacterial group from marine plankton. Nature 356:148-149.[CrossRef][Medline]
- Fuhrman, J. A., and R. T. Noble. 1995. Viruses and protists cause similar bacterial mortality in coastal seawaters. Limnol. Oceanogr. 40:1236-1242.
- Fuhrman, J. A., and C. C. Ouverney. 1998. Marine microbial diversity studied via 16S rRNA sequences: cloning results from coastal waters and counting of native Archaea with fluorescent single cell probes. Aquat. Ecol. 32:3-15.[CrossRef]
- Gasol, J. M. 1994. A framework for the assessment of top-down vs bottom-up control of heterotrophic nanoflagellate abundance. Mar. Ecol. Prog. Ser. 113:291-300.
- Gasol, J. M., M. Comerma, J. C. Garcia, J. Armengol, E. O. Casamayor, P. Kojecka, and K. Simek. 2002. A transplant experiment to identify the factors controlling bacterial abundance, activity, production, and community composition in a eutrophic canyon-shaped reservoir. Limnol. Oceanogr. 47:62-77.
- Gasol, J. M., and C. M. Duarte. 2000. Comparative analyses in aquatic microbial ecology: how far do they go? FEMS Microbiol. Ecol. 31:99-106.[CrossRef][Medline]
- Gasol, J. M., C. Pedros-Alio, and D. Vaqué. 2002. Regulation of bacterial assemblages in oligotrophic plankton systems: results from experimental and empirical approaches. Antonie van Leeuwenhoek 81:435-452.[CrossRef][Medline]
- Gil, A. I., V. R. Louis, I. N. G. Rivera, E. Lipp, A. Huq, C. F. Lanata, D. N. Taylor, E. Russek-Cohen, N. Choopun, R. B. Sack, and R. R. Colwell. 2004. Occurrence and distribution of Vibrio cholerae in the coastal environment of Peru. Environ. Microbiol. 6:699-706.[CrossRef][Medline]
- Giovannoni, S. J. 1991. The polymerase chain reaction, p. 177-203. In E. Stackebrandt and M. Goodfellow (ed.), Nucleic acid techniques in bacterial systematics. J. Wiley & Sons Ltd., West Sussex, UK.
- Giovannoni, S. J., T. B. Britschgi, C. L. Moyer, and K. G. Field. 1990. Genetic diversity in Sargasso Sea bacterioplankton. Nature 345:60-63.[CrossRef][Medline]
- Giovannoni, S. J., E. F. DeLong, G. J. Olsen, and N. R. Pace. 1988. Phylogenetic group-specific oligodeoxynucleotide probes for identification of single microbial cells. J. Bacteriol. 170:720-726.
[Abstract/Free Full Text] - Gleason, H. A. 1926. The individualistic concept of the plant association. Bull. Torrey Bot. Club 53:7-26.[CrossRef]
- Glöckner, F. O., R. Amann, A. Alfreider, J. Pernthaler, R. Psenner, K. Trebesius, and K.-H. Schleifer. 1996. An in situ hybridization protocol for detection and identification of planktonic bacteria. Syst. Appl. Microbiol. 19:403-406.
- Glöckner, F. O., B. M. Fuchs, and R. Amann. 1999. Bacterioplankton compositions of lakes and oceans: a first comparison based on fluorescence in situ hybridization. Appl. Environ. Microbiol. 65:3721-3726.
[Abstract/Free Full Text] - Glöckner, F. O., M. Kube, M. Bauer, H. Teeling, T. Lombardot, W. Ludwig, D. Gade, A. Beck, K. Borzym, K. Heitmann, R. Rabus, H. Schlesner, R. Amann, and R. Reinhardt. 2003. Complete genome sequence of the marine planctomycete Pirellula sp strain 1. Proc. Natl. Acad. Sci. USA 100:8298-8303.
[Abstract/Free Full Text] - Glöckner, F.-O., E. Zaichikov, N. Belkova, L. Denissova, J. Pernthaler, A. Pernthaler, and R. Amann. 2000. Comparative 16S rRNA analysis of lake bacterioplankton reveals globally distributed phylogenetic clusters including an abundant group of actinobacteria. Appl. Environ. Microbiol. 66:5053-5065.
[Abstract/Free Full Text] - Gobler, C. J., D. A. Hutchins, N. S. Fisher, E. M. Cosper, and S. A. Sanudo-Wilhelmy. 1997. Release and bioavailability of C, N, P, Se, and Fe following viral lysis of a marine chrysophyte. Limnol. Oceanogr. 42:1492-1504.
- González, J. M., R. P. Kiene, and M. A. Moran. 1999. Transformation of sulfur compounds by an abundant lineage of marine bacteria in the alpha-Subclass of the class Proteobacteria. Appl. Environ. Microbiol. 65:3810-3819.
[Abstract/Free Full Text] - González, J. M., F. Mayer, M. A. Moran, R. E. Hodson, and W. B. Whitman. 1997. Sagittula stellata gen. nov., sp. nov., a lignin-transforming bacterium from a coastal environment. Int. J. Syst. Bacteriol. 47:773-780.[Abstract]
- González, J. M., and M. A. Moran. 1997. Numerical dominance of a group of marine bacteria in the alpha-subclass of the class Proteobacteria in coastal seawater. Appl. Environ. Microbiol. 63:4237-4242.[Abstract]
- González, J. M., R. Simo, R. Massana, J. S. Covert, E. O. Casamayor, C. Pedros-Alio, and M. A. Moran. 2000. Bacterial community structure associated with a dimethylsulfoniopropionate-producing North Atlantic algal bloom. Appl. Environ. Microbiol. 66:4237-4246.
[Abstract/Free Full Text] - Gram, L., H. P. Grossart, A. Schlingloff, and T. Kiorboe. 2002. Possible quorum sensing in marine snow bacteria: production of acylated homoserine lactones by Roseobacter strains isolated from marine snow. Appl. Environ. Microbiol. 68:4111-4116.
[Abstract/Free Full Text] - Gray, N. D., R. Howarth, R. W. Pickup, J. G. Jones, and I. M. Head. 2000. Use of combined microautoradiography and fluorescence in situ hybridization to determine carbon metabolism in mixed natural communities of uncultured bacteria from the genus Achromatium. Appl. Environ. Microbiol. 66:4518-4522.
[Abstract/Free Full Text] - Grossart, H. P. 1999. Interactions between marine bacteria and axenic diatoms (Cylindrotheca fusiformis, Nitzschia laevis, and Thalassiosira weissflogii) incubated under various conditions in the lab. Aquat. Microb. Ecol. 19:1-11.
- Grossart, H. P., T. Kiorboe, K. Tang, and H. Ploug. 2003. Bacterial colonization of particles: growth and interactions. Appl. Environ. Microbiol. 69:3500-3509.
[Abstract/Free Full Text] - Grossart, H. P., and H. Ploug. 2000. Bacterial production and growth efficiencies: direct measurements on riverine aggregates. Limnol. Oceanogr. 45:436-445.
- Grossart, H. P., and H. Ploug. 2001. Microbial degradation of organic carbon and nitrogen on diatom aggregates. Limnol. Oceanogr. 46:267-277.
- Grossart, H. P., L. Riemann, and F. Azam. 2001. Bacterial motility in the sea and its ecological implications. Aquat. Microb. Ecol. 25:247-258.
- Grossart, H. P., G. F. Steward, J. Martinez, and F. Azam. 2000. A simple, rapid method for demonstrating bacterial flagella. Appl. Environ. Microbiol. 66:3632-3636.
[Abstract/Free Full Text] - Grossart, H.-P., and M. Simon. 1998. Bacterial colonization and microbial decomposition of limnetic organic aggregates (lake snow). Aquat. Microb. Ecol. 15:127-140.
- Grossart, H.-P., and M. Simon. 1993. Limnetic macroscopic organic aggregates (lake snow): Occurrence, characteristics, and microbial dynamics in Lake Constance. Limnol. Oceanogr. 38:532-546.
- Hagström, A., J. Pinhassi, and U. L. Zweifel. 2000. Biogeographical diversity among marine bacterioplankton. Aquat. Microb. Ecol. 21:231-244.
- Hagström, A., T. Pommier, F. Rohwer, K. Simu, W. Stolte, D. Svensson, and U. L. Zweifel. 2002. Use of 16S ribosomal DNA for delineation of marine bacterioplankton species. Appl. Environ. Microbiol. 68:3628-3633.
[Abstract/Free Full Text] - Hahn, M., and M. Pockl. 2005. Ecotypes of planktonic actinobacteria with identical 16S rRNA genes adapted to thermal niches in temperate, subtropical, and tropical freshwater habitats. Appl. Environ. Microbiol. 71:766-773.
[Abstract/Free Full Text] - Hahn, M. G. 2003. Isolation of strains belonging to the cosmopolitan Polynucleobacter necessarius cluster from freshwater habitats located in three climatic zones. Appl. Environ. Microbiol. 69:5248-5254.
[Abstract/Free Full Text] - Hahn, M. W., and M. G. Höfle. 1999. Flagellate predation on a bacterial model community: Interplay of size-selective grazing, specific bacterial cell size, and bacterial community composition. Appl. Environ. Microbiol. 65:4863-4872.
[Abstract/Free Full Text] - Hahn, M. W., and M. G. Höfle. 1998. Grazing pressure by a bacterivorous flagellate reverses the relative abundance of Comamonas acidovorans Px54 and Vibrio strain Cb5 in chemostat cocultures. Appl. Environ. Microbiol. 64:1910-1918.
[Abstract/Free Full Text] - Hahn, M. W., H. Lunsdorf, and L. Janke. 2004. Exopolymer production and microcolony formation by planktonic freshwater bacteria: defence against protistan grazing. Aquat. Microb. Ecol. 35:297-308.
- Hahn, M. W., H. Lünsdorf, Q. Wu, M. Schauer, M. G. Höfle, J. Boenigk, and P. Stadler. 2003. Isolation of novel ultramicrobacteria classified as actinobacteria from five freshwater habitats in Europe and Asia. Appl. Environ. Microbiol. 69:1442-1451.
[Abstract/Free Full Text] - Hahn, M. W., E. R. B. Moore, and M. G. Höfle. 1999. Bacterial filament formation, a defense mechanism against flagellate grazing, is growth rate controlled in bacteria of different phyla. Appl. Environ. Microbiol. 65:25-35.
[Abstract/Free Full Text] - Hamasaki, K., R. A. Long, and F. Azam. 2004. Individual cell growth rates of marine bacteria, measured by bromodeoxyuridine incorporation. Aquat. Microb. Ecol. 35:217-227.
- Heidelberg, J. F., K. B. Heidelberg, and R. R. Colwell. 2002. Seasonality of Chesapeake Bay bacterioplankton species. Appl. Environ. Microbiol. 68:5488-5497.
[Abstract/Free Full Text] - Heino, J., and J. Soininen. 2005. Assembly rules and community models for unicellular organisms: patterns in diatoms of boreal streams. Freshwater Biol. 50:567-577.[CrossRef]
- Herndl, G. J., G. Müller-Niklas, and J. Frick. 1993. Major role of ultraviolet B in controlling bacterioplankton growth in the surface layer of the ocean. Nature 361:717-719.[CrossRef]
- Herndl, G. J., T. Reinthaler, E. Teira, H. van Aken, C. Veth, A. Pernthaler, and J. Pernthaler. Contribution of Archaea to total prokaryotic production in the deep Atlantic Ocean. Appl. Environ. Microbiol. 71:2303-2309.
- Hiorns, W. D., B. A. Methe, S. A. Nierzwicki-Bauer, and J. P. Zehr. 1997. Bacterial diversity in Adirondack mountain lakes as revealed by 16S rRNA gene sequences. Appl. Environ. Microbiol. 63:2957-2960.[Abstract]
- Hobbie, J. E., R. J. Daley, and S. Jasper. 1977. Use of nuclepore filters for counting bacteria by fluorescence microscopy. Appl. Environ. Microbiol. 33:1225-1228.
[Abstract/Free Full Text] - Hollibaugh, J. T., N. Bano, and H. W. Ducklow. 2002. Widespread distribution in polar oceans of a 16S rRNA gene sequence with affinity to Nitrosospira-like ammonia-oxidizing bacteria. Appl. Environ. Microbiol. 68:1478-1484.
[Abstract/Free Full Text] - Hugenholtz, P., and T. Huber. 2003. Chimeric 16S rDNA sequences of diverse origin are accumulating in the public databases. Int. J. Syst. Evol. Bacteriol. 53:289-293.
[Abstract/Free Full Text] - Iriberri, J., I. Azua, A. Labirua-Iturburu, I. Artolozaga, and I. Barcina. 1994. Differential elimination of enteric bacteria by protists in a freshwater system. J. Appl. Bacteriol. 77:476-483.[Medline]
- Jannasch, H. W. 1974. Steady state and the chemostat in ecology. Limnol. Oceanogr. 19:716-720.
- Jaspers, E., K. Nauhaus, H. Cypionka, and J. Overmann. 2001. Multitude and temporal variability of ecological niches as indicated by the diversity of cultivated bacterioplankton. FEMS Microbiol. Ecol. 36:153-164.[CrossRef][Medline]
- Jaspers, E., and J. Overmann. 2004. Ecological significance of microdiversity: identical 16S rRNA gene sequences can be found in bacteria with highly divergent genomes and ecophysiologies. Appl. Environ. Microbiol. 70:4831-4839.
[Abstract/Free Full Text] - Jürgens, K., H. Arndt, and K. O. Rothhaupt. 1994. Zooplankton-mediated change of bacterial community structure. Microb. Ecol. 27:27-42.
- Jürgens, K., and H. Güde. 1994. The potential importance of grazing-resistant bacteria in planktonic systems. Mar. Ecol. Prog. Ser. 112:169-188.
- Jürgens, K., and C. Matz. 2002. Predation as a shaping force for the phenotypic and genotypic composition of planktonic bacteria. Antonie van Leeuwenhoek 81:413-434.[CrossRef][Medline]
- Jürgens, K., J. Pernthaler, S. Schalla, and R. Amann. 1999. Morphological and compositional changes in a planktonic bacterial community in response to enhanced protozoan grazing. Appl. Environ. Microbiol. 65:1241-1250.
[Abstract/Free Full Text] - Jürgens, K., O. Skibbe, and E. Jeppesen. 1999. Impact of metazooplankton on the composition and population dynamics of planktonic ciliates in a shallow, hypertrophic lake. Aquat. Microb. Ecol. 17:61-75.
- Jürgens, K., and G. Stolpe. 1995. Seasonal dynamics of crustacean zooplankton, heterotrophic nanoflagellates and bacteria in a shallow, eutrophic lake. Freshwater Biol. 33:27-38.[CrossRef]
- Kalmbach, S., W. Manz, B. Bendinger, and U. Szewzyk. 2000. In situ probing reveals Aquabacterium commune as a widespread and highly abundant bacterial species in drinking water biofilms. Water Res. 34:575-581.[CrossRef]
- Karner, M., E. F. DeLong, and D. M. Karl. 2001. Archaeal dominance in the mesopelagic zone of the Pacific Ocean. Nature 409:507-509.[CrossRef][Medline]
- Karner, M., and J. A. Fuhrman. 1997. Determination of active marine bacterioplankton: a comparison of universal 16S rRNA probes, autoradiography, and nucleoid staining. Appl. Environ. Microbiol. 63:1208-1213.[Abstract]
- Kemp, P. F., S. Lee, and J. LaRoche. 1993. Estimating the growth rate of slowly growing marine bacteria from RNA content. Appl. Environ. Microbiol. 59:2594-2601.
[Abstract/Free Full Text] - Kiene, R. P., L. J. Linn, J. González, M. A. Moran, and J. A. Bruton. 1999. Dimethylsulfoniopropionate and methanethiol are important precursors of methionine and protein-sulfur in marine bacterioplankton. Appl. Environ. Microbiol. 65:4549-4558.
[Abstract/Free Full Text] - Kiorboe, T., H. P. Grossart, H. Ploug, and K. Tang. 2002. Mechanisms and rates of bacterial colonization of sinking aggregates. Appl. Environ. Microbiol. 68:3996-4006.
[Abstract/Free Full Text] - Kiorboe, T., K. Tang, H. P. Grossart, and H. Ploug. 2003. Dynamics of microbial communities on marine snow aggregates: colonization, growth, detachment, and grazing mortality of attached bacteria. Appl. Environ. Microbiol. 69:3036-3047.
[Abstract/Free Full Text] - Kirchman, D., E. Knees, and R. Hodson. 1985. Leucine incorporation and its potential as a measure of protein-synthesis by bacteria in natural aquatic systems. Appl. Environ. Microbiol. 49:599-607.
[Abstract/Free Full Text] - Kirchman, D. L., A. I. Dittel, S. E. G. Findlay, and D. Fischer. 2004. Changes in bacterial activity and community structure in response to dissolved organic matter in the Hudson River, New York. Aquat. Microb. Ecol. 35:243-257.
- Kirchman, D. L., L. Y. Yu, and M. T. Cottrell. 2003. Diversity and abundance of uncultured Cytophaga-like bacteria in the Delaware Estuary. Appl. Environ. Microbiol. 69:6587-6596.
[Abstract/Free Full Text] - Kirchman, D. L., L. Y. Yu, B. M. Fuchs, and R. Amann. 2001. Structure of bacterial communities in aquatic systems as revealed by filter PCR. Aquat. Microb. Ecol. 26:13-22.
- Kitaguchi, H., N. Hiragushi, A. Mitsutani, M. Yamaguchi, and Y. Ishida. 2001. Isolation of an algicidal marine bacterium with activity against the harmful dinoflagellate Heterocapsa circularisquama (Dinophyceae). Phycologia 40:275-279.
- Langenheder, S., and K. Jürgens. 2001. Regulation of bacterial biomass and community structure by metazoan and protozoan predation. Limnol. Oceanogr. 46:121-134.
- Laurion, I., M. Ventura, J. Catalan, R. Psenner, and R. Sommaruga. 2000. Attenuation of ultraviolet radiation in mountain lakes: Factors controlling the among- and within-lake variability. Limnol. Oceanogr. 45:1274-1288.
- Lebaron, P., P. Catala, C. Fajon, F. Joux, J. Baudart, and L. Bernard. 1997. A new sensitive, whole-cell hybridization technique for detection of bacteria involving a biotinylated oligonucleotide probe targeting rRNA and tyramide signal amplification. Appl. Environ. Microbiol. 63:3274-3278.[Abstract]
- Lebaron, P., P. Servais, H. Agogue, C. Courties, and F. Joux. 2001. Does the high nucleic acid content of individual bacterial cells allow us to discriminate between active cells and inactive cells in aquatic systems? Appl. Environ. Microbiol. 67:1775-1782.
[Abstract/Free Full Text] - Lebaron, P., P. Servais, A. C. Baudoux, M. Bourrain, C. Courties, and N. Parthuisot. 2002. Variations of bacterial-specific activity with cell size and nucleic acid content assessed by flow cytometry. Aquat. Microb. Ecol. 28:131-140.
- Lebaron, P., P. Servais, M. Troussellier, C. Courties, J. Vives-Rego, G. Muyzer, L. Bernard, T. Guindulain, H. Schäfer, and E. Stackebrandt. 1999. Changes in bacterial community structure in seawater mesocosms differing in their nutrient status. Aquat. Microb. Ecol. 19:225-267.
- Ledyard, K. M., and J. W. H. Dacey. 1994. Dimethylsulfide production from dimethylsulfoniopropionate by a marine bacterium. Mar. Ecol. Prog. Ser. 110:95-103.
- Lee, N., P. Nielsen, K. Andreasen, S. Juretschko, J. Nielsen, K. Schleifer, and M. Wagner. 1999. Combination of fluorescent in situ hybridization and microautoradiography—a new tool for structure-function analyses in microbial ecology. Appl. Environ. Microbiol. 65:1289-1297.
[Abstract/Free Full Text] - Liesack, W., H. Weyland, and E. Stackebrandt. 1991. Potential risks of gene amplification by Pcr as determined By 16s rdna analysis of a mixed-culture of strict barophilic bacteria. Microb. Ecol. 21:191-198.
- Lindström, E. S., and A. K. Bergstrom. 2004. Influence of inlet bacteria on bacterioplankton assemblage composition in lakes of different hydraulic retention time. Limnol. Oceanogr. 49:125-136.
- Lindström, E. S., K. Vrede, and E. Leskinen. 2004. Response of a member of the Verrucomicrobia, among the dominating bacteria in a hypolimnion, to increased phosphorus availability. J. Plankton Res. 26:241-246.
[Abstract/Free Full Text] - Llobet-Brossa, E., R. Rossellò-Mora, and R. Amann. 1998. Microbial community composition of Wadden Sea sediments as revealed by fluorescence in situ hybridization. Appl. Environ. Microbiol. 64:2691-2696.
[Abstract/Free Full Text] - Loferer-Krössbacher, M., J. Klima, and R. Psenner. 1998. Determination of bacterial cell dry mass by transmission electron microscopy and densitometric image analysis. Appl. Environ. Microbiol. 64:688-694.
[Abstract/Free Full Text] - Long, R. A., and F. Azam. 2001. Antagonistic interactions among marine pelagic bacteria. Appl. Environ. Microbiol. 67:4975-4983.
[Abstract/Free Full Text] - Long, R. A., A. Qureshi, D. J. Faulkner, and F. Azam. 2003. 2-n-pentyl-4-quinolinol produced by a marine Alteromonas sp and its potential ecological and biogeochemical roles. Appl. Environ. Microbiol. 69:568-576.
[Abstract/Free Full Text] - Loy, A., M. Horn, and M. Wagner. 2003. probeBase: an online resource for rRNA-targeted oligonucleotide probes. Nucleic Acids Res. 31:514-516.
[Abstract/Free Full Text] - Ludwig, W., and H.-P. Klenk. 2001. Overview: a phylogenetic backbone and taxonomic framework for procayotic systematics, p. 49-65. In G. M. Garrety (ed.), Bergey's manual of systematic bacteriology, 2nd edition, vol. 1. Williams & Wilkins, New York, N.Y.
- Ludwig, W., O. Strunk, R. Westram, L. Richter, H. Meier, Yadhukumar, A. Buchner, T. Lai, S. Steppi, G. Jobb, W. Forster, I. Brettske, S. Gerber, A. W. Ginhart, O. Gross, S. Grumann, S. Hermann, R. Jost, A. Konig, T. Liss, R. Lussmann, M. May, B. Nonhoff, B. Reichel, R. Strehlow, A. Stamatakis, N. Stuckmann, A. Vilbig, M. Lenke, T. Ludwig, A. Bode, and K. H. Schleifer. 2004. ARB: a software environment for sequence data. Nucleic Acids Res. 32:1363-1371.
[Abstract/Free Full Text] - Macek, M., G. Carlos, P. Memije, and P. Ramirez. 1997. Ciliate Vibrio cholerae interactions within a microbial loop: an experimental study. Aquat. Microb. Ecol. 13:257-266.
- Mackenzie, C., M. Chidambaram, E. J. Sodergren, S. Kaplan, and G. M. Weinstock. 1995. DNA repair mutants of Rhodobacter sphaeroides. J. Bacteriol. 177:3027-3035.
[Abstract/Free Full Text] - Malin, G., W. H. Wilson, G. Bratbak, P. S. Liss, and N. H. Mann. 1998. Elevated production of dimethylsulfide resulting from viral infection of cultures of Phaeocystis pouchetii. Limnol. Oceanogr. 43:1389-1393.
- Malmstrom, R. R., M. T. Cottrell, H. Elifantz, and D. L. Kirchman. 2005. Biomass production and assimilation of dissolved organic matter by SAR11 bacteria in the northwest Atlantic Ocean. Appl. Environ. Microbiol. 71:2979-2986.
[Abstract/Free Full Text] - Malmstrom, R. R., R. P. Kiene, M. T. Cottrell, and D. L. Kirchman. 2004. Contribution of SAR11 bacteria to dissolved dimethylsulfoniopropionate and amino acid uptake in the North Atlantic ocean. Appl. Environ. Microbiol. 70:4129-4135.
[Abstract/Free Full Text] - Malmstrom, R. R., R. P. Kiene, and D. L. Kirchman. 2004. Identification and enumeration of bacteria assimilating dimethylsulfoniopropionate (DMSP) in the North Atlantic and Gulf of Mexico. Limnol. Oceanogr. 49:597-606.
- Manz, W., R. Amann, W. Ludwig, M. Wagner, and K.-H. Schleifer. 1992. Phylogenetic oligodeoxynucleotide probes for the major subclasses of Proteobacteria: problems and solutions. Syst. Appl. Microbiol. 15:593-600.
- Maranger, R., P. A. del Giorgio, and D. F. Bird. 2002. Accumulation of damaged bacteria and viruses in lake water exposed to solar radiation. Aquat. Microb. Ecol. 28:213-227.
- Massana, R., E. F. DeLong, and C. Pedros-Alio. 2001. A few cosmopolitan phylotypes dominate planktonic archaeal assemblages in widely different oceanic provinces. Appl. Environ. Microbiol. 66:1777-1787.[CrossRef]
- Massana, R., J. M. Gasol, P. K. Bjornsen, N. Blackburn, A. Hagström, S. Hietanen, B. H. Hygum, J. Kuparinen, and C. PedrosAlio. 1997. Measurement of bacterial size via image analysis of epifluorescence preparations: description of an inexpensive system and solutions to some of the most common problems. Sci. Mar. 61:397-407.
- Massana, R., and K. Jürgens. 2003. Composition and population dynamics of planktonic bacteria and bacterivorous flagellates in seawater chemostat cultures. Aquat. Microb. Ecol. 32:11-22.
- Massana, R., A. E. Murray, C. M. Preston, and E. F. Delong. 1997. Vertical distribution and phylogenetic characterization of marine planktonic Archaea in the Santa Barbara channel. Appl. Environ. Microbiol. 63:50-56.[Abstract]
- Massana, R., C. Pedros-Alio, E. O. Casamayor, and J. M. Gasol. 2001. Changes in marine bacterioplankton phylogenetic composition during incubations designed to measure biogeochemically significant parameters. Limnol. Oceanogr. 46:1181-1188.
- Matz, C., J. Boenigk, H. Arndt, and K. J.ürgens. 2002. Role of bacterial phenotypic traits in selective feeding of the heterotrophic nanoflagellate Spumella sp. Aquat. Microb. Ecol. 27:137-148.
- McCandliss, R. R., S. E. Jones, M. Hearn, R. Latter, and C. F. Jago. 2002. Dynamics of suspended particles in coastal waters (southern North Sea) during a spring bloom. J. Sea Res. 47:285-302.[CrossRef]
- Methé, B. A., W. D. Hiorns, and J. P. Zehr. 1998. Contrasts between marine and freshwater bacterial community composition: Analyses of communities in Lake George and six other Adirondack lakes. Limnol. Oceanogr. 43:368-374.
- Middelboe, M., A. Hagström, N. Blackburn, B. Sinn, U. Fischer, N. H. Borch, J. Pinhassi, K. Simu, and M. G. Lorenz. 2001. Effects of bacteriophages on the population dynamics of four strains of pelagic marine bacteria. Microb. Ecol. 42:395-406.[CrossRef][Medline]
- Middelboe, M., N. O. G. Jorgensen, and N. Kroer. 1996. Effects of viruses on nutrient turnover and growth efficiency of noninfected marine bacterioplankton. Appl. Environ. Microbiol. 62:1991-1997.[Abstract]
- Moeseneder, M. M., J. M. Arrieta, G. Muyzer, C. Winter, and G. J. Herndl. 1999. Optimization of terminal-restriction fragment length polymorphism analysis for complex marine bacterioplankton communities and comparison with denaturing gradient gel electrophoresis. Appl. Environ. Microbiol. 65:3518-3525.
[Abstract/Free Full Text] - Moeseneder, M. M., C. Winter, J. M. Arrieta, and G. J. Herndl. 2001. Terminal-restriction fragment length polymorphism (T-RFLP) screening of a marine archaeal clone library to determine the different phylotypes. J. Microbiol. Methods 44:159-172.[CrossRef][Medline]
- Moeseneder, M. M., C. Winter, and G. J. Herndl. 2001. Horizontal and vertical complexity of attached and free-living bacteria of the eastern Mediterranean Sea, determined by 16S rDNA and 16S rRNA fingerprints. Limnol. Oceanogr. 46:95-107.
- Monger, B. C., and M. R. Landry. 1991. Prey size dependency of grazing by freeliving marine flagellates. Mar. Ecol. Prog. Ser. 74:239-248.
- Morita, R. Y. 1997. Bacteria in oligotrophic environments: starvation-survival lifestyle, vol. 1. Chapman Hall, New York, N.Y.
- Morris, R. M., M. S. Rappe, S. A. Connon, K. L. Vergin, W. A. Siebold, C. A. Carlson, and S. J. Giovannoni. 2002. SAR11 clade dominates ocean surface bacterioplankton communities. Nature 420:806-810.[CrossRef][Medline]
- Morris, R. M., M. S. Rappe, E. Urbach, S. A. Connon, and S. J. Giovannoni. 2004. Prevalence of the chloroflexi-related SAR202 bacterioplankton cluster throughout the mesopelagic zone and deep ocean. Appl. Environ. Microbiol. 70:2836-2842.
[Abstract/Free Full Text] - Mourino-Perez, R. R., A. Z. Worden, and F. Azam. 2003. Growth of Vibrio cholerae O1 in red tide waters off California. Appl. Environ. Microbiol. 69:6923-6931.
[Abstract/Free Full Text] - Murray, A. E., A. Blakis, R. Massana, S. Strawzewski, U. Passow, A. Alldredge, and E. F. DeLong. 1999. A time series assessment of planktonic archaeal variability in the Santa Barbara Channel. Aquat. Microb. Ecol. 20:129-145.
- Murray, A. E., C. M. Preston, R. Massana, L. T. Taylor, A. Blakis, K. Wu, and E. F. Delong. 1998. Seasonal and spatial variability of bacterial and archaeal assemblages in the coastal waters near Anvers Island, Antarctica. Appl. Environ. Microbiol. 64:2585-2595.
[Abstract/Free Full Text] - Muyzer, G., E. C. de Waal, and A. G. Uitterlinden. 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol. 59:695-700.
[Abstract/Free Full Text] - Muyzer, G., A. Teske, C. O. Wirsen, and H. W. Jannasch. 1995. Phylogenetic relationship of Thiomicrospira species and their identification in deep-sea hydrothermal vent samples by denaturing gradient gel electrophoresis of 16S rDNA fragments. Arch. Microbiol. 164:165-172.[CrossRef][Medline]
- Nedoma, J., A. Strojsova, J. Vrba, J. Komarkova, and K. Simek. 2003. Extracellular phosphatase activity of natural plankton studied with ELF97 phosphate: fluorescence quantification and labelling kinetics. Environ. Microbiol. 5:462-472.[CrossRef][Medline]
- Nielsen, J. L., M. A. de Muro, and P. H. Nielsen. 2003. Evaluation of the redox dye 5-cyano-2,3-tolyl-tetrazolium chloride for activity studies by simultaneous use of microautoradiography and fluorescence in situ hybridization. Appl. Environ. Microbiol. 69:641-643.
[Abstract/Free Full Text] - Norland, S. 1993. The relationship between biomass and volume of bacteria, p. 303-307. In P. Kemp, B. F. Sherr, E. B. Sherr, and J. Cole (ed.), Handbook of methods in aquatic microbial ecology. Lewis, Boca Raton, Fla.
- Orphan, V. J., C. H. House, K. U. Hinrichs, K. D. McKeegan, and E. F. DeLong. 2001. Methane-consuming archaea revealed by directly coupled isotopic and phylogenetic analysis. Science 293:484-487.
[Abstract/Free Full Text] - Ouverney, C. C., and J. A. Fuhrman. 1999. Combined microautoradiography-16S rRNA probe technique for determination of radioisotope uptake by specific microbial cell types in situ. Appl. Environ. Microbiol. 65:1746-1752.
[Abstract/Free Full Text] - Ouverney, C. C., and J. A. Fuhrman. 1997. Increase in fluorescence intensity of 16S rRNA in situ hybridization in natural samples treated with chloramphenicol. Appl. Environ. Microbiol. 63:2735-2740.[Abstract]
- Ouverney, C. C., and J. A. Fuhrman. 2000. Marine planktonic archaea take up amino acids. Appl. Environ. Microbiol. 66:4829-4833.
[Abstract/Free Full Text] - Ovreas, L., D. Bourne, R. A. Sandaa, E. O. Casamayor, S. Benlloch, V. Goddard, G. Smerdon, M. Heldal, and T. F. Thingstad. 2003. Response of bacterial and viral communities to nutrient manipulations in seawater mesocosms. Aquat. Microb. Ecol. 31:109-121.
- Pace, M. L., and J. J. Cole. 1994. Comparative and experimental approaches to top-down and bottom-up regulation of bacteria. Microb. Ecol. 28:181-193.[CrossRef]
- Page, K. A., S. A. Connon, and S. J. Giovannoni. 2004. Representative freshwater bacterioplankton isolated from Crater Lake, Oregon. Appl. Environ. Microbiol. 70:6542-6550.
[Abstract/Free Full Text] - Partensky, F., W. R. Hess, and D. Vaulot. 1999. Prochlorococcus, a marine photosynthetic prokaryote of global significance. Microbiol. Mol. Biol. Rev. 63:106-127.
[Abstract/Free Full Text] - Passow, U. 2002. Transparent exopolymer particles (TEP) in aquatic environments. Prog. Oceanogr. 55:287-333.[CrossRef]
- Pedros-Alio, C., and S. Y. Newell. 1989. Microautoradiographic study of thymidine uptake in brackish waters around Sapelo Island, Georgia, USA Mar. Ecol. Prog. Ser. 55:83-94.
- Pernthaler, A., and R. Amann. 2004. Simultaneous fluorescence in situ hybridization of mRNA and rRNA in environmental bacteria. Appl. Environ. Microbiol. 70:5526-5533.
- Pernthaler, A., and J. Pernthaler. 2005. Diurnal variation of cell proliferation in three bacterial taxa from coastal North Sea waters. Appl. Environ. Microbiol. 71:4638-4644.
[Abstract/Free Full Text] - Pernthaler, A., J. Pernthaler, and R. Amann. 2002. Fluorescence in situ hybridization and catalyzed reporter deposition for the identification of marine bacteria. Appl. Environ. Microbiol. 68:3094-3101.
[Abstract/Free Full Text] - Pernthaler, A., J. Pernthaler, and R. Amann. 2004. Sensitive multicolour fluorescence in situ hybridization for the identification of environmental organisms, p. 711-726. In G. A. Kowalchuk, De Bruijn, F.J., I. M. Head, A. D. L. Akkermans, and J. D. van Elsas (ed.), Molecular microbial ecology manual, 2nd ed. Kluwer Academic Publishers, Dordecht, The Netherlands.
- Pernthaler, A., J. Pernthaler, H. Eilers, and R. Amann. 2001. Growth patterns of two marine isolates: adaptations to substrate patchiness? Appl. Environ. Microbiol. 67:4077-4083.
[Abstract/Free Full Text] - Pernthaler, A., J. Pernthaler, M. Schattenhofer, and R. Amann. 2002. Identification of DNA-synthesizing bacterial cells in coastal North Sea plankton. Appl. Environ. Microbiol. 68:5728-5736.
[Abstract/Free Full Text] - Pernthaler, A., C. M. Preston, J. Pernthaler, E. F. DeLong, and R. Amann. 2002. A comparison of fluorescently labeled oligonucleotide and polynucleotide probes for the detection of pelagic marine bacteria and archaea. Appl. Environ. Microbiol. 68:661-668.
[Abstract/Free Full Text] - Pernthaler, J. 2005. Predation on procaryotes in the water column and its ecological implications. Nat. Rev. Microbiol. 3:537-546.[CrossRef][Medline]
- Pernthaler, J., A. Alfreider, T. Posch, S. Andreatta, and R. Psenner. 1997. In situ classification and image cytometry of pelagic bacteria from a high mountain lake (Gossenköllesee, Austria). Appl. Environ. Microbiol. 63:4778-4783.[Abstract]
- Pernthaler, J., F. O. Glöckner, W. Schönhuber, and R. Amann. 2001. Fluorescence in situ hybridization (FISH) with rRNA-targeted oligonucleotide probes, p. 207-226. In J. H. Paul (ed.), Methods in microbiology, vol. 30. Academic Press, San Diego, Calif.[CrossRef]
- Pernthaler, J., F. O. Glöckner, S. Unterholzner, A. Alfreider, R. Psenner, and R. Amann. 1998. Seasonal community and population dynamics of pelagic Bacteria and Archaea in a high mountain lake. Appl. Environ. Microbiol. 64:4299-4306.
[Abstract/Free Full Text] - Pernthaler, J., A. Pernthaler, and R. Amann. 2003. Automated enumeration of groups of marine picoplankton after fluorescence in situ hybridization. Appl. Environ. Microbiol. 69:2631-2637.
[Abstract/Free Full Text] - Pernthaler, J., T. Posch, K. Simek, J. Vrba, R. Amann, and R. Psenner. 1997. Contrasting bacterial strategies to coexist with a flagellate predator in an experimental microbial assemblage. Appl. Environ. Microbiol. 63:596-601.[Abstract]
- Pernthaler, J., T. Posch, K. Simek, J. Vrba, A. Pernthaler, F. O. Glöckner, U. Nübel, R. Psenner, and R. Amann. 2001. Predator-specific enrichment of actinobacteria from a cosmopolitan freshwater clade in mixed continuous culture. Appl. Environ. Microbiol. 67:2145-2155.
[Abstract/Free Full Text] - Pernthaler, J., B. Sattler, K. Simek, A. Schwarzenbacher, and R. Psenner. 1996. Top-down effects on the size-biomass distribution of a freshwater bacterioplankton community. Aquat. Microb. Ecol. 10:255-263.
- Pernthaler, J., E. Zöllner, F. Warnecke, and K. J.ürgens. 2004. Blooms of filamentous bacteria in a mesotrophic lake: identity and potential controlling mechanisms. Appl. Environ. Microbiol. 70:6272-6281.
[Abstract/Free Full Text] - Pinhassi, J., and T. Berman. 2003. Differential growth response of colony-forming alpha- and gammaproteobacteria in dilution culture and nutrient addition experiments from Lake Kinneret (Israel), the eastern Mediterranean Sea, and the Gulf of Eilat. Appl. Environ. Microbiol. 69:199-211.
[Abstract/Free Full Text] - Pinhassi, J., M. M. Sala, H. Havskum, F. Peters, O. Guadayol, A. Malits, and C. L. Marrase. 2004. Changes in bacterioplankton composition under different phytoplankton regimens. Appl. Environ. Microbiol. 70:6753-6766.
[Abstract/Free Full Text] - Pinhassi, J., A. Winding, S. J. Binnerup, U. L. Zweifel, B. Riemann, and A. Hagström. 2003. Spatial variability in bacterioplankton community composition at the Skagerrak-Kattegat Front. Mar. Ecol. Prog. Ser. 255:1-13.
- Pinhassi, J., and A. Hagström. 2000. Seasonal succession in marine bacterioplankton. Aquat. Microb. Ecol. 21:245-256.
- Pinhassi, J., U. L. Zweifel, and A. Hagström. 1997. Dominant marine bacterioplankton species found among colony-forming bacteria. Appl. Environ. Microbiol. 63:3359-3366.[Abstract]
- Ploug, H., H. P. Grossart, F. Azam, and B. B. Jorgensen. 1999. Photosynthesis, respiration, and carbon turnover in sinking marine snow from surface waters of Southern California Bight: implications for the carbon cycle in the ocean. Mar. Ecol. Prog. Ser. 179:1-11.
- Ploug, H., M. Kühl, B. Buchholz Cleven, and B. B. J.örgensen. 1997. Anoxic aggregates: An ephemeral phenomenon in the pelagic environment? Aquat. Microb. Ecol. 13:285-294.
- Porter, K. G., and Y. S. Feig. 1980. The use of DAPI for identifying and counting aquatic microflora. Limnol. Oceanogr. 25:943-948.
- Posch, T., J. Jezbera, J. Vrba, K. Simek, J. Pernthaler, S. Andreatta, and B. Sonntag. 2001. Size selective feeding in Cyclidium glaucoma (Ciliophora, Scuticociliatida) and its effects on bacterial community structure: A study from a continuous cultivation system. Microb. Ecol. 42:217-227.[CrossRef][Medline]
- Posch, T., M. Loferer-Krössbacher, G. Gao, A. Alfreider, J. Pernthaler, and R. Psenner. 2001. Precision of bacterioplankton biomass determination: a comparison of two fluorescent dyes, and of allometric and linear volume-to-carbon conversion factors. Aquat. Microb. Ecol. 25:55-63.
- Posch, T., K. Simek, J. Vrba, S. Pernthaler, J. Nedoma, B. Sattler, B. Sonntag, and R. Psenner. 1999. Predator-induced changes of bacterial size-structure and productivity studied on an experimental microbial community. Aquat. Microb. Ecol. 18:235-246.
- Psenner, R., and R. Sommaruga. 1992. Are rapid changes in bacterial biomass caused by shifts from top-down to bottom-up control. Limnol. Oceanogr. 37:1092-1100.
- Radajewski, S., P. Ineson, N. R. Parekh, and J. C. Murrell. 2000. Stable-isotope probing as a tool in microbial ecology. Nature 403:646-649.[CrossRef][Medline]
- Ramsing, N. B., H. Fossing, G. Ferdelman Timothy, F. Andersen, and B. Thamdrup. 1996. Distribution of bacterial populations in a stratified Fjord (Mariager Fjord, Denmark) quantified by in situ hybridization and related to chemical gradients in the water column. Appl. Environ. Microbiol. 62:1391-1404.[Abstract]
- Rappe, M. S., S. A. Connon, K. L. Vergin, and S. J. Giovannoni. 2002. Cultivation of the ubiquitous SAR11 marine bacterioplankton clade. Nature 418:630-633.[CrossRef][Medline]
- Rappe, M. S., and S. J. Giovannoni. 2003. The uncultured microbial majority. Annu. Rev. Microbiol. 57:369-394.[CrossRef][Medline]
- Rappe, M. S., D. A. Gordon, K. L. Vergin, and S. J. Giovannoni. 1999. Phylogeny of actinobacteria small subunit (SSU) rRNA gene clones recovered from marine bacterioplankton. Syst. Appl. Microbiol. 22:106-112.
- Rappe, M. S., K. Vergin, and S. J. Giovannoni. 2000. Phylogenetic comparisons of a coastal bacterioplankton community with its counterparts in open ocean and freshwater systems. FEMS Microbiol. Ecol. 33:219-232.[CrossRef][Medline]
- Reysenbach, A. L., L. J. Giver, G. S. Wickham, and N. R. Pace. 1992. Differential amplification of ribosomal RNA genes by polymerase chain reaction. Appl. Environ. Microbiol. 58:3417-3418.
[Abstract/Free Full Text] - Riemann, L., and F. Azam. 2002. Widespread N-acetyl-D-glucosamine uptake among pelagic marine bacteria and its ecological implications. Appl. Environ. Microbiol. 68:5554-5562.
[Abstract/Free Full Text] - Riemann, L., G. F. Steward, and F. Azam. 2000. Dynamics of bacterial community composition and activity during a mesocosm diatom bloom. Appl. Environ. Microbiol. 66:578-587.
[Abstract/Free Full Text] - Rizzoli, R., N. M. Maraldi, A. Galanzi, N. Zini, M. Falconi, M. Vitale, and G. Mazzotti. 1988. High-sensitivity detection of DNA-synthesis by immunolocalization of bromodeoxyuridine. Institute of Physics Conference Series 93:551-552.
- Rocap, G., F. W. Larimer, J. Lamerdin, S. Malfatti, P. Chain, N. A. Ahlgren, A. Arellano, M. Coleman, L. Hauser, W. R. Hess, Z. I. Johnson, M. Land, D. Lindell, A. F. Post, W. Regala, M. Shah, S. L. Shaw, C. Steglich, M. B. Sullivan, C. S. Ting, A. Tolonen, E. A. Webb, E. R. Zinser, and S. W. Chisholm. 2003. Genome divergence in two Prochlorococcus ecotypes reflects oceanic niche differentiation. Nature 424:1042-1047.[CrossRef][Medline]
- Rodriguez, G., D. Phipps, K. Ishiguro, and H. Ridgway. 1992. Use of a fluorescent redox probe for direct visualization of actively respiring bacteria. Appl. Environ. Microbiol. 58:1801-1808.
[Abstract/Free Full Text] - Rossello-Mora, R., and R. Amann. 2001. The species concept for prokaryotes. FEMS Microbiol. Rev. 25:39-67.[CrossRef][Medline]
- Rossello-Mora, R. A., M. Wagner, R. Amann, and K.-H. Schleifer. 1995. The abundance of Zoogloea ramigera in sewage treatment plants. Appl. Environ. Microbiol. 61:702-707.[Abstract]
- Salcher, M. M., J. Pernthaler, R. Psenner, and T. Posch. Succession of bacterial grazing defense mechanisms against protistan predators in an experimental microbial community. Aquat. Microb. Ecol. 38:215-229.
- Sanders, R. W., D. A. Caron, and U. G. Berninger. 1992. Relationship between bacteria and heterotrophic nanoplankton in marine and freshwaters: an inter-ecosystem comparison. Mar. Ecol. Prog. Ser. 86:1-14.
- Schäfer, H., B. Abbas, H. Witte, and G. Muyzer. 2002. Genetic diversity of ‘satellite’ bacteria present in cultures of marine diatoms. FEMS Microbiol. Ecol. 42:25-35.[CrossRef]
- Schäfer, H., L. Bernard, C. Courties, P. Lebaron, P. Servais, R. Pukall, E. Stackebrandt, M. Troussellier, T. Guindulain, J. Vives-Rego, and G. Muyzer. 2001. Microbial community dynamics in Mediterranean nutrient-enriched seawater. FEMS Microbiol. Ecol. 34:243-253.[CrossRef][Medline]
- Schäfer, H., P. Servais, and G. Muyzer. 2000. Successional changes in the genetic diversity of a marine assemblage during confinement. Arch. Microbiol. 173:138-145.[CrossRef][Medline]
- Schauer, M., and M. Hahn. 2005. Diversity and phylogenetic affiliations of morphologically conspicuous large filamentous bacteria occurring in the pelagic zones of a broad spectrum of freshwater habitats. Appl. Environ. Microbiol. 71:1931-1940.
[Abstract/Free Full Text] - Schink, B. 2000. Structure and function relationships in natural microbial communities. FEMS Microbiol. Rev. 24:553-553.[CrossRef][Medline]
- Schramm, A., L. H. Larsen, N. P. Revsbech, N. B. Ramsing, R. Amann, and K.-H. Schleifer. 1996. Structure and function of a nitrifying Biofilm as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 62:4641-4647.[Abstract]
- Schulze, R., S. Spring, R. Amann, I. Huber, W. Ludwig, K.-H. Schleifer, and P. Kämpfer. 1999. Genotypic diversity of Acidovorax strains isolated from activated sludge and description of Acidovorax defuvii sp. nov. Syst. Appl. Microbiol. 22:205-214.[Medline]
- Schut, F., M. Jansen, T. M. P. Gomes, J. C. Gottschal, W. Harder, and R. A. Prins. 1995. Substrate uptake and utilization by a marine ultramicrobacterium. Microbiology 141:351-361.[Abstract]
- Schwalbach, M. S., I. Hewson, and J. A. Fuhrman. 2004. Viral effects on bacterial community composition in marine plankton microcosms. Aquat. Microb. Ecol. 34:117-127.
- Schweitzer, B., I. Huber, R. Amann, W. Ludwig, and M. Simon. 2001. Alpha- and betaproteobacteria control the consumption and release of amino acids on lake snow aggregates. Appl. Environ. Microbiol. 67:632-645.
[Abstract/Free Full Text] - Sekar, R., B. Fuchs, R. Amann, and J. Pernthaler. 2004. Flow Sorting of marine bacterioplankton after fluorescence in situ hybridization. Appl. Environ. Microbiol. 70:6210-6219.
[Abstract/Free Full Text] - Sekar, R., A. Pernthaler, J. Pernthaler, F. Warnecke, T. Posch, and R. Amann. 2003. An improved protocol for the quantification of freshwater actinobacteria by fluorescence in situ hybridization. Appl. Environ. Microbiol. 69:2928-2935.
[Abstract/Free Full Text] - Sekiguchi, H., M. Watanabe, T. Nakahara, B. H. Xu, and H. Uchiyama. 2002. Succession of bacterial community structure along the Changjiang River determined by denaturing gradient gel electrophoresis and clone library analysis. Appl. Environ. Microbiol. 68:5142-5150.
[Abstract/Free Full Text] - Selje, N., and M. Simon. 2003. Composition and dynamics of particle-associated and free-living bacterial communities in the Weser estuary, Germany. Aquat. Microb. Ecol. 30:221-237.
- Selje, N., M. Simon, and T. Brinkhoff. 2004. A newly discovered Roseobacter cluster in temperate and polar oceans. Nature 427:445-448.[CrossRef][Medline]
- Servais, P., C. Courties, P. Lebaron, and M. Troussellier. 1999. Coupling bacterial activity measurements with cell sorting by flow cytometry. Microb. Ecol. 38:180-189.[CrossRef][Medline]
- Sherr, B., E. Sherr, and P. del Giorgio. 2001. Enumeration of total and highly active bacteria. Methods Microbiol. 30:129-159.[CrossRef]
- Sherr, B. F., E. B. Sherr, and J. McDaniel. 1992. Effect of protistan grazing on the frequency of dividing cells in bacterioplankton assemblages. Appl. Environ. Microbiol. 58:4371-4378.
- Sherr, B. F., E. B. Sherr, and C. Pedros Alio. 1989. Simultaneous measurement of bacterioplankton production and protozoan bacterivory in estuarine water. Mar. Ecol. Prog. Ser. 54:209-219.
- Sherr, E. B., and B. F. Sherr. 1994. Bacterivory and herbivory: key roles of phagotrophic protists in pelagic food webs. Microb. Ecol. 28:223-235.[CrossRef]
- Sherr, E. B., B. F. Sherr, and L. J. Albright. 1987. Bacteria—link or sink. Science 235:88-88.
- Shikano, S., L. S. Luckinbill, and Y. Kurihara. 1990. Changes of traits in a bacterial population associated with protozoal predation. Microb. Ecol. 20:75-84.
- Sieracki, M. E., P. W. Johnson, and J. M. Sieburth. 1985. Detection, enumeration, and sizing of planktonic bacteria by image analyzed epifluorescence microscopy. Appl. Environ. Microbiol. 49:799-810.
[Abstract/Free Full Text] - Simek, K., J. Armengol, M. Comerma, J. C. Garcia, P. Kojecka, J. Nedoma, and J. Hejzlar. 2001. Changes in the epilimnetic bacterial community composition, production, and protist-induced mortality along the longitudinal axis of a highly eutrophic reservoir. Microb. Ecol. 42:359-371.[CrossRef][Medline]
- Simek, K., and T. H. Chrzanowski. 1992. Direct and indirect evidence of size-selective grazing on pelagic bacteria by freshwater nanoflagellates. Appl. Environ. Microbiol. 58:3715-3720.
[Abstract/Free Full Text] - Simek, K., J. Nedoma, J. Pernthaler, T. Posch, and J. R. Dolan. 2002. Altering the balance between bacterial production and protistan bacterivory triggers shifts in freshwater bacterial community composition. Antonie van Leeuwenhoek 81:453-463.[CrossRef][Medline]
- Simek, K., J. Pernthaler, M. Weinbauer, K. Hornak, J. Dolan, J. Nedoma, M. Masin, and R. Amann. 2001. Changes in bacterial community composition, dynamics and viral mortality rates associated with enhanced flagellate grazing in a mesoeutrophic reservoir. Appl. Environ. Microbiol. 67:2723-2733.
[Abstract/Free Full Text] - Simek, K., J. Vrba, J. Pernthaler, T. Posch, P. Hartman, J. Nedoma, and R. Psenner. 1997. Morphological and compositional shifts in an experimental bacterial community influenced by protists with contrasting feeding modes. Appl. Environ. Microbiol. 63:587-595.[Abstract]
- Simon, M., F. O. Glöckner, and R. Amann. 1999. Different community structure and temperature optima of heterotrophic picoplankton in various regions of the Southern Ocean. Aquat. Microb. Ecol. 18:275-284.
- Simon, M., H. P. Grossart, B. Schweitzer, and H. Ploug. 2002. Microbial ecology of organic aggregates in aquatic ecosystems. Aquat. Microb. Ecol. 28:175-211.
- Simu, K., and A. Hagström. 2004. Oligotrophic bacterioplankton with a novel single-cell life strategy. Appl. Environ. Microbiol. 70:2445-2451.
[Abstract/Free Full Text] - Skovhus, T. L., N. B. Ramsing, C. Holmstrom, S. Kjelleberg, and I. Dahllof. 2004. Real-time quantitative PCR for assessment of abundance of Pseudoalteromonas species in marine samples. Appl. Environ. Microbiol. 70:2373-2382.
[Abstract/Free Full Text] - Snaidr, J., B. Fuchs, G. Wallner, M. Wagner, K.-H. Schleifer, and R. Amann. 1999. Phylogeny and in situ identification of a morphologically conspicuous bacterium, Candidatus Magnospira bakii, present at very low frequency in activated sludge. Environ. Microbiol. 1:125-135.[CrossRef][Medline]
- Sommaruga, R., I. Obernosterer, G. J. Herndl, and R. Psenner. 1997. Inhibitory effect of solar radiation on thymidine and leucine incorporation by freshwater and marine bacterioplankton. Appl. Environ. Microbiol. 63:4178-4184.[Abstract]
- Sommaruga, R., and R. Psenner. 1995. Permanent presence of grazing-resistant bacteria in a hypertrophic lake. Appl. Environ. Microbiol. 61:3457-3459.[Abstract]
- Stach, J. E. M., L. A. Maldondando, A. C. Ward, M. Goodfellow, and A. T. Bull. 2003. New primers for the class Actinobacteria: application to marine and terrestrial environments. Environ. Microbiol. 5:828-841.[CrossRef][Medline]
- Stoderegger, K. E., and G. J. Herndl. 2002. Distribution of capsulated bacterioplankton in the North Atlantic and North Sea. Microb. Ecol. 44:154-163.[CrossRef][Medline]
- Straskrabová, V. 1983. The effect of substrate shock on populations of starving aquatic bacteria. J. Appl. Bacteriol. 54:217-224.
- Stürmeyer, H., J. Overmann, H. D. Babenzien, and H. Cypionka. 1998. Ecophysiological and phylogenetic studies of Nevskia ramosa in pure culture. Appl. Environ. Microbiol. 64:1890-1894.
[Abstract/Free Full Text] - Sunamura, M., Y. Higashi, C. Miyako, J. Ishibashi, and A. Maruyama. 2004. Two bactetia phylotypes are predominant in the Suiyo Seamount hydrothermal plume. Appl. Environ. Microbiol. 70:1190-1198.
[Abstract/Free Full Text] - Suzuki, M., M. S. Rappe, and S. J. Giovannoni. 1998. Kinetic bias in estimates of coastal picoplankton community structure obtained by measurements of small-subunit rRNA gene PCR amplicon length heterogeneity. Appl. Environ. Microbiol. 64:4522-4529.
[Abstract/Free Full Text] - Suzuki, M. T. 1999. Effect of protistan bacterivory on coastal bacterioplankton diversity. Aquat. Microb. Ecol. 20:261-272.
- Suzuki, M. T., and S. J. Giovannoni. 1996. Bias caused by template annealing in the amplification of mixtures of 16S rRNA genes by PCR. Appl. Environ. Microbiol. 62:625-630.[Abstract]
- Suzuki, M. T., C. M. Preston, F. P. Chavez, and E. F. DeLong. 2001. Quantitative mapping of bacterioplankton populations in seawater: field tests across an upwelling plume in Monterey Bay. Aquat. Microb. Ecol. 24:117-127.
- Teira, E., T. Reinthaler, A. Pernthaler, J. Pernthaler, and G. J. Herndl. 2004. Combining catalyzed reporter deposition-fluorescence in situ hybridization and microautoradiography to detect substrate utilization by bacteria and archaea in the deep ocean. Appl. Environ. Microbiol. 70:4411-4414.
[Abstract/Free Full Text] - Teske, A., E. Alm, J. M. Regan, S. Toze, B. E. Rittmann, and D. A. Stahl. 1994. Evolutionary relationships among ammonia- and nitrite-oxidizing bacteria. J. Bacteriol. 176:6623-6630.
[Abstract/Free Full Text] - Thingstad, T. F. 2000. Elements of a theory for the mechanisms controlling abundance, diversity, and biogeochemical role of lytic bacterial viruses in aquatic systems. Limnol. Oceanogr. 45:1320-1328.
- Thurman, E. 1985. Organic geochemistry of natural waters. Martinus Nijhoff Publishers, Dordrecht, The Netherlands.
- Trebesius, K., R. Amann, W. Ludwig, K. Mühlegger, and K.-H. Schleifer. 1994. Identification of whole fixed bacterial cells with nonradioactive 23S rRNA-targeted polynucleotide probes. Appl. Environ. Microbiol. 60:3228-3235.
[Abstract/Free Full Text] - Tsai, Y.-L., and B. H. Olson. 1991. Rapid method for direct extration of DNA from soil and sediments. Appl. Environ. Microbiol. 57:1070-1074.
[Abstract/Free Full Text] - Urbach, E., K. L. Vergin, and S. J. Giovannoni. 1999. Immunochemical detection and isolation of DNA from metabolically active bacteria. Appl. Environ. Microbiol. 65:1207-1213.
[Abstract/Free Full Text] - Urbach, E., K. L. Vergin, L. Young, A. Morse, G. L. Larson, and S. J. Giovannoni. 2001. Unusual bacterioplankton community structure in ultra- oligotrophic Crater Lake. Limnol. Oceanogr. 46:557-572.
- van Hannen, E. J., G. Zwart, M. P. van Agterveld, H. J. Gons, J. Ebert, and H. J. Laanbroek. 1999. Changes in bacterial and eukaryotic community structure after mass lysis of filamentous cyanobacteria associated with viruses. Appl. Environ. Microbiol. 65:795-801.
[Abstract/Free Full Text] - Venter, J. C., K. Remington, J. F. Heidelberg, A. L. Halpern, D. Rusch, J. A. Eisen, D. Y. Wu, I. Paulsen, K. E. Nelson, W. Nelson, D. E. Fouts, S. Levy, A. H. Knap, M. W. Lomas, K. Nealson, O. White, J. Peterson, J. Hoffman, R. Parsons, H. Baden-Tillson, C. Pfannkoch, Y. H. Rogers, and H. O. Smith. 2004. Environmental genome shotgun sequencing of the Sargasso Sea. Science 304:66-74.
[Abstract/Free Full Text] - Vila, M., R. Simo, R. P. Kiene, J. Pinhassi, J. A. González, M. A. Moran, and C. Pedros-Alio. 2004. Use of microautoradiography combined with fluorescence in situ hybridization to determine dimethylsulfoniopropionate incorporation by marine bacterioplankton taxa. Appl. Environ. Microbiol. 70:4648-4657.
[Abstract/Free Full Text] - Vrba, J., C. Callier, T. Bittl, K. Simek, R. Bertoni, P. Filandr, P. Hartman, J. Hejzlar, M. Macek, and J. Nedoma. 2004. Are bacteria the major producers of extracellular glycolytic enzymes in aquatic environments? Int. Rev. Hydrobiol. 89:102-117.[CrossRef]
- Wagner, M., A. Loy, R. Nogueira, U. Purkhold, N. Lee, and H. Daims. 2002. Microbial community composition and function in wastewater treatment plants. Antonie van Leeuwenhoek 81:665-680.[CrossRef][Medline]
- Warnecke, F., J. Pernthaler, and R. Amann. 2004. Actinobacterial 16S rRNA genes from freshwater habitats cluster in four distinct lineages. Environ. Microbiol. 6:242-253.[CrossRef][Medline]
- Warnecke, F., R. Sommaruga, J. S. Hofer, and J. Pernthaler. Abundances, identity, and growth state of actinobacteria in mountain lakes of different transparencies. Appl. Environ. Microbiol. 71:5561-5569.
- Weinbauer, M. G. 2004. Ecology of prokaryotic viruses. FEMS Microbiol. Rev. 28:127-181.[CrossRef][Medline]
- Weinbauer, M. G., and M. G. Höfle. 1998. Distribution and life strategies of two bacterial populations in a eutrophic lake. Appl. Environ. Microbiol. 64:3776-3783.
[Abstract/Free Full Text] - Weisse, T. 1991. The microbial food web and its sensitivity to eutrophication and contaminant enrichment—a cross-system overview. Int. Rev. Ges. Hydrobiol. 76:327-338.
- West, N. J., W. A. Schönhuber, N. J. Fuller, R. I. Amann, R. Rippka, A. F. Post, and D. J. Scanlan. 2001. Closely related Prochlorococcus genotypes show remarkably different depth distributions in two oceanic regions as revealed by in situ hybridization using 16S rRNA-targeted oligonucleotides. Microbiology 147:1731-1744.
[Abstract/Free Full Text] - Whitaker, R. J., D. W. Grogan, and J. W. Taylor. 2003. Geographic barriers isolate endemic populations of hyperthermophilic archaea. Science 301:976-978.
[Abstract/Free Full Text] - Winogradsky, S. 1888. Beiträge zur Morphologie und Physiologie der Bakterien, Heft 1: Zur Morphologie und Physiologie der Schwefelbakterien. Arthur Felix, Leipzig, Germany.
- Winter, C., A. Smit, G. J. Herndl, and M. G. Weinbauer. 2004. Impact of virioplankton on archaeal and bacterial community richness as assessed in seawater batch cultures. Appl. Environ. Microbiol. 70:804-813.
[Abstract/Free Full Text] - Wohl, D. L., S. Arora, and J. R. Gladstone. 2004. Functional redundancy supports biodiversity and ecosystem function in a closed and constant environment. Ecology. 85:1534-1540.
- Wommack, K. E., and R. R. Colwell. 2000. Virioplankton: viruses in aquatic ecosystems. Microbiol. Mol. Biol. Rev. 64:69-114.
[Abstract/Free Full Text] - Worden, A. Z., S. W. Chisholm, and B. J. Binder. 2000. In situ hybridization of Prochlorococcus and Synechococcus (marine cyanobacteria) spp. with rRNA-targeted peptide nucleic acid probes. Appl. Environ. Microbiol. 66:284-289.
[Abstract/Free Full Text] - Wuchter, C., S. Schouten, H. T. S. Boschker, and J. S. S. Damste. 2003. Bicarbonate uptake by marine Crenarchaeota. FEMS Microbiol. Lett. 219:203-207.[CrossRef][Medline]
- Zimmermann, R., R. Iturriaga, and J. Beckerbirck. 1978. Simultaneous determination of total number of aquatic bacteria and number thereof involved in respiration. Appl. Environ. Microbiol. 36:926-935.
[Abstract/Free Full Text] - Zubkov, M. V., B. M. Fuchs, S. D. Archer, R. P. Kiene, R. Amann, and P. A. Burkill. 2001. Linking the composition of bacterioplankton to rapid turnover of dissolved dimethylsulfoniopropionate in an algal bloom in the North Sea. Environ. Microbiol. 3:304-311.[CrossRef][Medline]
- Zubkov, M. V., B. M. Fuchs, S. D. Archer, R. P. Kiene, R. Amann, and P. H. Burkill. 2002. Rapid turnover of dissolved DMS and DMSP by defined bacterioplankton communities in the stratified euphotic zone of the North Sea. Deep-Sea Res. Part Ii Top. Studies Oceanogr. 49:3017-3038.
- Zubkov, M. V., B. M. Fuchs, P. H. Burkill, and R. Amann. 2001. Comparison of cellular and biomass specific activities of dominant bacterioplankton groups in stratified waters of the Celtic Sea. Appl. Environ. Microbiol. 67:5210-5218.
[Abstract/Free Full Text] - Zubkov, M. V., B. M. Fuchs, H. Eilers, P. H. Burkill, and R. Amann. 1999. Determination of total protein content of bacterial cells by SYPRO staining and flow cytometry. Appl. Environ. Microbiol. 65:3251-3257.
[Abstract/Free Full Text] - Zubkov, M. V., B. M. Fuchs, G. A. Tarran, P. H. Burkill, and R. Amann. 2003. High rate of uptake of organic nitrogen compounds by Prochlorococcus cyanobacteria as a key to their dominance in oligotrophic oceanic waters. Appl. Environ. Microbiol. 69:1299-1304.
[Abstract/Free Full Text] - Zwart, G., B. C. Crump, M. Agterveld, F. Hagen, and S. K. Han. 2002. Typical freshwater bacteria: an analysis of available 16S rRNA gene sequences from plankton of lakes and rivers. Aquat. Microb. Ecol. 28:141-155.
- Zwart, G., W. Hiorns, B. Methe, M. Van Agterveld, R. Huismans, S. Nold, J. Zehr, and H. Laanbroek. 1998. Nearly identical 16S rRNA sequences recovered from lakes in North America and Europe indicate the existence of clades of globally distributed freshwater bacteria. Syst. Appl. Microbiol. 21:546-556.[Medline]
- Zwart, G., R. Huismans, M. P. van Agterveld, Y. Van de Peer, P. De Rijk, H. Eenhoorn, G. Muyzer, E. J. van Hannen, H. J. Gons, and H. J. Laanbroek. 1998. Divergent members of the bacterial division Verrucomicrobiales in a temperate freshwater lake. FEMS Microbiol. Ecol. 25:159-169.[CrossRef]
- Zwart, G., E. J. van Hannen, M. P. Kamst-van Agterveld, K. Van der Gucht, E. S. Lindstrom, J. Van Wichelen, T. Lauridsen, B. C. Crump, S. K. Han, and S. Declerck. 2003. Rapid screening for freshwater bacterial groups by using reverse line blot hybridization. Appl. Environ. Microbiol. 69:5875-5883.
[Abstract/Free Full Text]
Microbiology and Molecular Biology Reviews, September 2005, p. 440-461, Vol. 69, No. 3
1092-2172/05/$08.00+0 doi:10.1128/MMBR.69.3.440-461.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
This article has been cited by other articles:
-
Dimitriu, P. A., Pinkart, H. C., Peyton, B. M., Mormile, M. R.
(2008). Spatial and Temporal Patterns in the Microbial Diversity of a Meromictic Soda Lake in Washington State. Appl. Environ. Microbiol.
74: 4877-4888
[Abstract] [Full Text] -
Hunt, D. E., David, L. A., Gevers, D., Preheim, S. P., Alm, E. J., Polz, M. F.
(2008). Resource Partitioning and Sympatric Differentiation Among Closely Related Bacterioplankton. Science
320: 1081-1085
[Abstract] [Full Text] -
Giddens, S. R., Jackson, R. W., Moon, C. D., Jacobs, M. A., Zhang, X.-X., Gehrig, S. M., Rainey, P. B.
(2007). Mutational activation of niche-specific genes provides insight into regulatory networks and bacterial function in a complex environment. Proc. Natl. Acad. Sci. USA
104: 18247-18252
[Abstract] [Full Text] -
Polz, M. F, Hunt, D. E, Preheim, S. P, Weinreich, D. M
(2006). Patterns and mechanisms of genetic and phenotypic differentiation in marine microbes. Phil Trans R Soc B
361: 2009-2021
[Abstract] [Full Text]
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Copyright © 2009 by the American Society for Microbiology. For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»