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Applied and Environmental Microbiology, June 1999, p. 2478-2484, Vol. 65, No. 6
Department of Microbial
Ecology,1 and Department of Food Web
Studies,2 Centre for Limnology, Netherlands
Institute of Ecology, 3600 BG Maarssen, The Netherlands
Received 9 December 1998/Accepted 18 March 1999
Correlations between the biomass of phytoplankton and the biomass
of bacteria and between the biomass of bacteria and the biomass of
protozoans suggest that there is coupling between these compartments of
the "microbial loop." To investigate this coupling on the species
level, bacteria and protozoans from untreated lake water inocula were
allowed to grow on detritus of the green alga Ankistrodesmus
falcatus or the cyanobacterium Oscillatoria limnetica in continuous-flow systems for 1 month. Denaturing gradient gel electrophoresis (DGGE) of the 16S and 18S rRNA genes was used to
monitor the development of the bacterial community structure and the
eukaryotic community structure, respectively. Nonmetric multidimensional scaling of the DGGE profiles revealed the changes in
the microbial community structure. This analysis showed that significantly different bacterial communities developed on the green
algal detritus and on the cyanobacterial detritus. Although similar
results were obtained for the eukaryotic communities, the differences
were not significant. Hence, our findings indicate that the origin of
detritus can affect the structure of at least the bacterial community.
A phylogenetic analysis of 20 18S ribosomal DNA clones that were
isolated from the continuous cultures revealed that many sequences were
related to the sequences of bacterivorous protozoans (members of the
Ciliophora, Rhizopoda, Amoeba, and Kinetoplastida). One clone grouped
in a recently established clade whose previously described members are
all parasites. The affiliations of about 20% of the clones could not
be determined.
In aquatic environments,
photosynthetically produced organic carbon is decomposed by
heterotrophic bacteria, which in turn are consumed by heterotrophic
protozoans. Correlations between phytoplankton biomass and
heterotrophic bacterial biomass (10, 12, 41) and between
heterotrophic bacterial biomass and protozoan biomass (8, 9, 11,
28) suggest that there is coupling between the compartments of
the so-called microbial loop (4). However, most descriptions
of such relationships have been based on bulk measurements of the
compartments, and whether there are correlations at the species level
is an intriguing question. For example, is organic carbon derived from
different phytoplankton species decomposed by different species of
specialized bacteria? Or is protozoan species composition governed by
bacterial species composition or just by the concentration of edible
food particles? Although numerous studies have shown that bacteria have
different rates of growth on natural carbon sources (6, 7, 16, 21, 32, 33) and that heterotrophic protozoans selectively graze on
larger bacteria (15, 29, 31), little is known about whether the development of aquatic microbial communities depends on various organic carbon sources. This lack of information is attributable to the
difficulty of determining the presence of microbial species due to the
fact that the majority of these microorganisms cannot be grown with the
current cultivation methods. Molecular techniques can be used to assess
genetic composition without culturing all members of a community. To
investigate the potential dependence of microbial community structure
on the source of detritus in lakes, we used denaturing gradient gel
electrophoresis (DGGE). We determined the bacterial community structure
and the eukaryotic community structure (22, 35) for
communities that developed on detritus derived from the green alga
Ankistrodesmus falcatus or on detritus derived from the
cyanobacterium Oscillatoria limnetica in specially designed
two-stage continuous-flow systems (34). Since DGGE
does not provide quantitative results (38), we converted the
band patterns to reflect whether particular sequence types were
present or absent. Our qualitative measures of microbial community
structure were analyzed by nonmetric multidimensional scaling (NMDS)
and a statistical method used for comparisons of the DGGE patterns.
Special attention was paid to the eukaryotic microbial community. Since
most heterotrophic protozoans are bacterivorous, we wanted to determine
whether the carbon source used by the bacteria also influenced the
community structure of the protozoans.
Experimental setup.
Nonaxenic A. falcatus (Centre
for Limnology strain E01) and Oscillatoria cf.
limnetica (Centre for Limnology strain MR1) were grown under
light-limiting conditions in modified Guillard medium (34)
in the first stages (volume, 2 liters) of two continuous-flow systems
(designated the Alga system and the Cyano system) until the steady
state was reached (dilution rate, 0.36 day
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Detritus-Dependent Development of the Microbial Community in
an Experimental System: Qualitative Analysis by Denaturing
Gradient Gel Electrophoresis
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
1). The
two-stage continuous-flow system has been described previously (34). The cultures were grown at a constant temperature
(20°C) at pH 8.0 ± 0.2 with illumination by type TLE 32W/33
circular fluorescent lamps (Philips). The inner sides of the lamps
facing the culture vessels were shielded with aluminum foil to decrease the light intensity. The light intensity for the
Ankistrodesmus cultures (Alga system) was manipulated so
that the chlorophyll a (Chla) concentrations in
the two systems were approximately the same. Samples (10 ml per assay)
were removed every 24 h and used to determine the concentrations
of Chla and total suspended solids (TSS).
2; mean depth, 3 m) with a short water retention
time (~1 week). This lake water was used as an inoculum for bacteria
and heterotrophic eukaryotes and was chosen because both cyanobacteria
and green algae are present in Lake Ketelmeer (36). If the
origin of detritus governs microbial community structure, the lake
water inoculum should have contained microorganisms that were
specialized for decomposing these carbon sources. The second stages
received UV-C-killed phytoplankton from the first stages at a dilution
rate of 0.30 day1. The UV-C intensities needed to kill
O. limnetica and A. falcatus were 5 and 18 W
· m2, respectively. The procedures used to assess
lethal UV-C intensities have been described previously (34).
For 1 month, samples were removed and used for Chla, TSS,
and community structure analyses (DGGE and 18S rRNA sequence determination).
Chla and TSS. The Chla and TSS assays used have been described previously (34). Duplicate measurements were obtained with each assay.
DNA extraction, PCR, and DGGE. DNA was released from cells by mechanical force (bead beating), phenol extraction, and ethanol precipitation (42). PCR primers for the V2 region were used for amplification of the 16S rRNA gene. The PCR primers used were F357GC (5'-CGCCCGCCGCGCCCCGCGCCCGGCCCGCCGCCCCCGCCCCCCTACGGGAGGCAGCAG-3'), which contains a GC-rich clamp and is specific for most members of the Bacteria, and R518 (5'-ATTACCGCGGCTGCTGG-3'), which is specific for most members of the Bacteria, Archaea, and Eucarya (22). Each PCR amplification was performed in a 50-µl reaction mixture containing approximately 100 ng of template DNA, 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 0.01% (wt/vol) gelatin, 1.5 mM MgCl2, each primer at a concentration of 0.5 µM, each deoxynucleotide at a concentration of 200 µM, 400 ng of bovine serum albumin, and 2.5 U of Taq DNA polymerase (Boehringer, Mannheim, Germany). PCR cycling was performed with a Perkin-Elmer model 480 thermocycler. The temperature cycling conditions were as follows: preincubation at 94°C for 5 min, followed by 25 cycles consisting of 94°C for 1 min, the annealing temperature (TA) for 1 min, and 72°C for 1 min. In the first 20 cycles the TA was decreased by 1°C stepwise each two cycles, from 65°C in the first cycle to 56°C in the 20th cycle. In the last five cycles the TA was 55°C. The 25 cycles were followed by 5 min of incubation at 72°C. The primers used for amplification of the 18S rRNA gene were F1427GC (5'-CGCCCGCCGCGCCCCGCGCCCGGCCCGCCGCCCCCGCCCCTCTGTGATGCCCTTAGATGTTCTGGG-3') and R1616 (5'-GCGGTGTGTACAAAGGGCAGGG-3'). Both of these primers are specific for eukaryotic aquatic microorganisms (35). The temperature cycling conditions were as follows: preincubation at 94°C for 5 min, followed by 25 cycles consisting of 94°C for 1 min, 52°C for 1 min, and 72°C for 1 min and then a final extension step consisting of 72°C for 5 min. The reaction conditions were the same as described above. DGGE was performed as described previously (35, 42). Briefly, PCR products of similar sizes were separated on a 1.5-mm-thick vertical gel containing 8% (wt/vol) polyacrylamide (acrylamide/bisacrylamide ratio, 37.5:1) and a linear gradient consisting of the denaturants urea and formamide; the concentration of the denaturants increased from 35% at the top of the gel to 55% at the bottom for separation of the 16S ribosomal DNA (rDNA) fragment and from 30 to 55% for separation of the 18S rDNA fragment (100% denaturant was defined as 7 M urea and 40% [vol/vol] formamide). Equal amounts of PCR products were applied to the DGGE gel. The concentrations of PCR products were estimated by separating the products on 2.0% agarose gels, staining them with ethidium bromide (see below), and analyzing digitized images with ImageQuant software (Molecular Dynamics Ltd., Kemsing, England). A 50-µl portion of the sample containing the smallest amount of PCR product was loaded onto the DGGE gel; all other samples were loaded in amounts relative to this sample. Electrophoresis was performed at 60°C with a buffer containing 40 mM Tris, 40 mM acetic acid, and 1 mM EDTA (pH 7.6) (0.5× TAE buffer), and 75 V was applied to the submerged gel for 16 h. Nucleic acids were visualized by staining the gel for 1 h in 0.5× TAE buffer containing 0.5 mg of ethidium bromide per liter and then destaining it for 5 min in demineralized water and photographing it with a charge-coupled device camera (The Imager; Appligene, Illkirch, France). Digitized images were inverted by using the Photostyler software (Aldus Corporation, Seattle, Wash.). The contrast and gray balance of the entire image were adjusted to reduce the background.
Clone library construction. To estimate the eukaryotic species present in the continuous-flow systems, the DNA extracted from three samples (Alga 1 stage on day 23, Alga 2 stage on day 23, and Cyano 1 stage on day 29) were mixed and used for clone library construction. 18S rDNA clones that were nearly full length were generated by using the Eucarya-specific primers E4 (5'-CTGGTTGATTCTGCCAGT-3') and E1628 (5'-CGACGGGCGGTGTGTA-3') (the numbers in the designations indicate Saccharomyces cerevisiae sequence positions). Each PCR amplification was performed in a 50-µl reaction mixture containing approximately 100 ng of template DNA, 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 0.01% (wt/vol) gelatin, 1.5 mM MgCl2, each primer at a concentration of 0.5 µM, each deoxynucleotide at a concentration of 200 µM, 400 ng of bovine serum albumin, and 2.5 U of Taq DNA polymerase (Boehringer). PCR cycling was performed with the Perkin-Elmer model 480 thermocycler programmed as follows: denaturation at 94°C for 5 min, followed by 25 cycles consisting of 94°C for 1 min, 54°C for 1 min, and 72°C for 1 min and a final extension step consisting of 68°C for 10 min. The cloning and sequencing procedures used have been described previously (42). A total of 120 clones were examined by DGGE. Only clones whose DGGE band positions were different were sequenced. The sequences were determined in two directions by using the following Texas red-labeled primers: M13 forward (5'-TGTAAAACGACGGCCAGT-3'), M13 reverse (5'-GAAACAGCTATGACCATG-3'), E382 forward (5'-CGGAGAGGGAGCCTGAG-3'), E565 reverse (5'-ATTACCGCGGCTGCTGG-3'), E1128 forward (5'-AAACTTAAAGGAATTGACG-3'), and E1179 reverse (5'-CCCGTGTTGAGTCAAATT-3'), (the numbers in the designations indicate S. cerevisiae sequence positions).
Phylogenetic analysis. The 18S rDNA sequences obtained were compared with sequences obtained from GenBank/EMBL by using BLAST (1). The sequences with the highest levels of similarity were used as reference sequences for alignment. The alignment was based on secondary structure and was constructed by using the Dedicated Comparative Sequence Editor (13). Phylogenetic trees were constructed by using only unambiguously aligned sequence positions and maximum-likelihood analysis (PAUP*, test version 4.0d59; David L. Swofford, Laboratory of Molecular Systematics, Smithsonian Institution, Washington, D.C.). Each analysis was performed five times. Nucleotide frequencies and transition-to-transversion ratios were estimated from the data. Nucleotide substitution rates were assumed to follow a gamma distribution with a shape parameter of 0.5 and setting according to the HKY model (18). "Tree-bisection-reconnection" was used as a swapping algorithm. Maximum-parsimony bootstrap analysis (1,000 replicates) was used to assess the robustness of the trees.
Data analysis.
The DGGE patterns were converted to a binary
(01) matrix (35). A Nei-Li distance matrix was calculated
from the binary data (23). This distance matrix was analyzed
by using the NMDS package from the Statistica software package
(StatSoft, Inc., Tulsa, Okla.). This procedure presented the data in a
Euclidean plane such that very similar values were plotted close
together. The resulting graphical representation (NMDS map) was much
easier to interpret than the original table of distances was. When it
was applied to DGGE data, the NMDS map showed every band pattern (a
reflection of the community structure at a particular point in time) as
one point, and relative changes in community structure could be
visualized and interpreted by connecting consecutive points (37,
38). The statistical significance of the differences in community
structure due to the source of detritus was determined by using the
Nei-Li distances and comparing the communities only at the same moment in time. Two communities fed with different types of detritus were
considered to be different at a given moment in time if their Nei-Li
distance was significantly greater than the Nei-Li distances between
replicate communities. For each of the two sources of detritus we used
two replicates and made six observations at different times. On the
first sampling day (day 3) the distances between replicates were
exceptionally small compared to the distances on later days, indicating
that there was a lack of independence from the shared inoculum.
Therefore, these distances were not included in the calculations. Thus,
a total of 10 distances between replicates were used for the bacterial
and eukaryotic community to calculate an average distance between
replicates (
). Each
distance between communities growing on different sources of detritus
was tested against this distribution and was considered significant if
Disalga-cyano,t > 
+ (T · Stddis-repl.), where
Disalga-cyano,t is the distance between two
communities fed with algal or cyanobacterial detritus at time
t, T is the Student t value
(n = 9; P = 0.05), and
Stddis-repl. is the standard deviation of
.
Nucleotide sequence accession numbers. The eukaryotic clone sequences have been deposited in the EMBL database under accession no. AJ130849 to AJ130869.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Decomposition of detritus. The Chla and TSS concentrations in both first stages (Alga system and Cyano system) are shown in Fig. 1. Although the first stages of both flow systems were at a steady state at the beginning of the experiment, in the Alga system the Chla concentration increased after 3 days, while the TSS concentration was constant for the first 15 days. In the Cyano system the Chla level was constant throughout the experiment, while the TSS concentration decreased during the first 15 days. As a result, the two systems had different Chla concentrations after day 15, but their TSS concentrations were about the same. On a dry weight basis, the second stages of the two flow systems received approximately the same amounts of UV-killed biomass after 2 weeks.
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Microbial community structure. In the 16S rDNA DGGE analysis (Fig. 2A) 54 different bands were detected, while 58 different bands were detected in the 18S rDNA DGGE analysis (Fig. 2B). The relative changes in the structures of the bacterial and eukaryotic communities, as visualized by NMDS, are shown in Fig. 2C and D. Three days after the experiment started, the bacterial communities (Fig. 2C) growing on different types of detritus were still quite similar. A similar observation was made for the eukaryotic communities (Fig. 2D). A clear divergence between the bacterial communities grown on different types of detritus occurred on day 7. This trend persisted, and on day 17 the differences became significantly greater than the differences between the replicates (P < 0.05). Although the structures of the eukaryotic communities also changed, the trend was not as clear as the trend in the bacterial communities, and the differences between the replicates were as great as the differences between the treatments (green algal detritus versus cyanobacterial detritus).
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, Alga 1 stage; 
, Alga 2 stage; 
, Cyano 1 stage; 
,
Cyano 2 stage. The lines connect the community structures of replicate
stages. The numbers indicate the numbers of days from the beginning of
the experiment. Boldface numbers indicate significant differences
(P < 0.05) in community structures between different
sources of detritus. The white bands in panel B, lane c, indicate the
band positions for the 18S rDNA clones (Table Eukaryotic phylogeny. Of the 120 clones analyzed, 20 were found to have different DGGE bands. The sequences of these 20 clones were determined and phylogenetically analyzed by maximum-likelihood and parsimony methods (Fig. 3), and the frequency with which a band was found at the same position as the clones in the different stages of the two systems is shown in Table 1. Since 58 different band positions were identified on the 18S rDNA DGGE gels, our clone library accounted for at most one-third of the total eukaryotic diversity in the two systems. Of the 20 clones, 5 belonged to the Ciliophora (Fig. 3A), 3 belonged to the Rhizopoda (Fig. 3C), 3 belonged to the Metazoa (Fig. 3D), 1 belonged to the Drips (Fig. 3D), 1 belonged to the Amoeba (Fig. 3B), 1 belonged to the fungi (Fig. 3B), and 1 belonged to the Kinetoplastida (Fig. 3E), and the affiliations of 4 clones were not determined. Three of the sequences formed a unique terminal cluster with the fungi as the closest relatives (Fig. 3B).
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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We performed an experiment in which we investigated whether the origin of detritus can be a factor that governs microbial community structure. Although we found differences between replicates of the bacterial community growing on the same type of detritus, the effect of the carbon source (green algal detritus versus cyanobacterial detritus) was significantly greater (Fig. 2). In contrast to the bacterial community, the eukaryotic community developed rather chaotically; i.e., the differences between detritus sources were obscured by differences between replicates. This might be interpreted as due to poor reproducibility of the microcosm experiments and to the fact that development of microbial communities at the physical scale which we used is not determined solely by culture conditions, such as light, temperature, and nutrients. However, it is too soon for general conclusions to be made.
The abundance of a sequence was not included in our measurements of community structure since we used only the presence or absence of a DGGE band. Therefore, the changes in the community structure which we measured reflected only changes in species composition, not changes in species abundance. In addition, changes in the community structure, as detected by NMDS analysis, were influenced by the total number of bands in a DGGE pattern and by the number of bands shared by DGGE patterns. Differences in community structure could, therefore, have originated from differences in species diversity (the number of different DGGE bands obtained for two communities) or from differences in species richness (the total number of DGGE bands per community). In general, the bacterial richness of the Alga systems was somewhat greater than the bacterial richness of the Cyano systems. However, the differences in community structure that depended on the source of detritus were largely due to differences in bacterial diversity. The eukaryotic richness fluctuated greatly between sampling days and in all second stages. Thus, differences in the eukaryotic community structure between duplicates and between sources of detritus originated from variations in both eukaryotic richness and diversity.
Our results show that the origin of detritus can affect the structure of the microbial community, at least the bacterial community in our systems. Variations in the chemical composition of phytoplankton (25, 27, 30) and in bacterial substrate utilization may explain the effects on the bacterial community. Although some authors have described chemosensory feeding of bacterivorous protozoans (40), there is very strong evidence that size-selective feeding by protozoans occurs (15, 29, 31). If protozoan prey selection is based solely on size, one might not expect that detritus would have a strong effect on eukaryotic community structure unless the size spectrum of the bacterial community also changes due to the origin of the detritus.
Although we observed no significant differences between the eukaryotic
communities growing on the different types of detritus, it was
interesting to gain insight into the phylogenetic positions of the
protozoan species developing in the flow systems, especially since
eukaryotic microorganisms are largely neglected in molecular ecology.
Since we constructed our clone library from three samples obtained on
two sampling dates, this library does not represent the complete
eukaryotic community. We obtained 20 different clones, while
eukaryotic DGGE produced 58 different bands. Thus, our library accounts for one-third of the DGGE-detected eukaryotic diversity. Surprisingly, the clone library did not include A. falcatus.
Apparently, the high UV-C intensity (18 W · m2)
needed to kill this alga destroyed its DNA. The phylogenetic position
of the 18S rDNA clones was inferred by maximum-likelihood analysis
(Fig. 3). Replicate analyses (n = 5) always resulted in
the same phylogenetic tree. Since the second stages were kept in the
dark, there was no algal and or cyanobacterial growth, and bacterivory
or uptake of detritus or dissolved organic matter was the only possible
mode of feeding. Not surprisingly, most of the sequences found
were related to sequences of bacterivorous eukaryotes (members of the
Ciliophora, Amoeba Rhizopoda, and Kinetoplastida). Microscopic
observations confirmed that these bacterivorous eukaryotes were
present. However, we did not identify species. Two sequences (LKM80 and
LKM85) were closely related to the sequence of Brachionus plicatilis, a rotifer species capable of feeding on all members of
the microbial loop (3). One sequence (LKM88) was closely related to the sequence of a Chaetonotus sp. (Gastrotricha).
Since we did not prefilter the lake water inoculum, the metazoan
grazers were not eliminated and could develop in the second stages of the flow systems. Three sequences (LKM11, LKM15, and LKM46) were not
related to any of the eukaryotic sequences in the EMBL database and
formed a unique terminal clade supported by a moderately strong parsimony bootstrap value. The closest relatives of these sequences were the fungi (Fig. 3B). We provisionally designated this cluster LKM11 after the first clone. Another sequence, LKM118, was not related
to any previously described eukaryotic sequence. Phylogenetic analysis
placed the LKM118 sequence just below the "crown" of the eukaryotic
tree, although this position was very unstable (data not shown).
One sequence (LKM51) is particularly interesting. This sequence grouped in a recently established phylogenetic clade, the DRIPs clade, which was provisionally named after its first members (26). All of the sequences currently in this cluster are from protistan parasites of fish, crustaceans, and amphibians, and LKM51 exhibited a very high level of similarity (97.6%) with the "Prototheca richardsi" sequence. The latter sequence is from an agent that inhibits growth of amphibian larvae and was previously classified as a unicellular unpigmented alga (5). "P. richardsi" is the only DRIPs species whose free-living stage has been observed. None of the other DRIPs members have been cultured in vivo, and therefore the complete life histories of these parasites remain unclear. Apparently, our culture method supported growth of a free-living DRIPs species and may be used for further investigations of the life histories of these protistan parasites. Since we did not identify the eukaryotic microorganisms by microscopy, we have no information concerning the morphology of the species represented by sequence LKM51.
The resistance of most microbial species to cultivation has hampered the study of microbial community structure for many years. Recently introduced molecular techniques circumvent the problems and have been used increasingly to solve problems in microbial ecology. rDNA sequence information is now starting to reveal patterns and governing forces in natural microbial community structure (24). We used DGGE of the 16S and 18S rRNA genes to show that the origin of detritus can be a governing force in microbial community structure. Although many workers have used detritus as a natural carbon source to investigate bacterial production (17, 19, 39), to our knowledge we are the first researchers to describe detritus-dependent development of the microbial loop. Even though our study should be considered a survey and was carried out in small-scaled continuous-flow systems, the occurrence and subsequent decline of single-species phytoplankton blooms (2, 20) may result in organic carbon releases comparable to carbon releases in our flow system. The effects described above can be expected to take place in natural systems.
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ACKNOWLEDGMENTS |
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We thank P. Schouten, H. Uittenhout, and G. M. van Hannen for construction of continuous culture boxes and UV-C devices and T. Beebe and M. Ragan for sharing the "P. richardsi" sequence before publication.
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FOOTNOTES |
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* Corresponding author. Mailing address: Department of Microbial Ecology, Centre for Limnology, Netherlands Institute of Ecology, P.O. Box 1299, 3600 BG Maarssen, The Netherlands. Phone: 31 (0)294 239300. Fax: 31 (0)294 232224. E-mail: vanhannen{at}cl.nioo.knaw.nl.
Publication no. 2525 of the Centre for Limnology, Netherlands
Institute of Ecology, Maarssen, The Netherlands.
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REFERENCES |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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| 1. | Altschul, S. F., W. Gish, W. Miller, E. W. Myers, and D. J. Lipman. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403-410[Medline]. |
| 2. | Amon, R. M. W., and R. Benner. 1994. Rapid cycling of high-molecular weight dissolved organic matter in the ocean. Nature 369:549-552. |
| 3. |
Arndt, H.
1993.
Rotifers as predators on components of the microbial web (bacteria, heterotrophic flagellates, ciliates) a review.
Hydrobiologia
255/256:231-246.
|
| 4. | 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. |
| 5. | Beebee, T. Personal communication. |
| 6. | Bell, W. H. 1980. Bacterial utilization of algal extracellular products. 1. The kinetic approach. Limnol. Oceanogr. 25:1007-1020. |
| 7. | Bell, W. H., and E. Sakshaug. 1980. Bacterial utilization of algal extracellular products. 2. A kinetic study of natural populations. Limnol. Oceanogr. 25:1021-1033. |
| 8. | Berninger, U.-G., B. J. Finlay, and P. Kuuppo-Leinikki. 1991. Protozoan control of bacterial abundances in freshwater. Limnol. Oceanogr. 36:139-147. |
| 9. | Berninger, U.-G., S. A. Wickham, and B. J. Finlay. 1993. Trophic coupling within the microbial food web: a study with fine temporal resolution in a eutrophic freshwater ecosystem. Freshwater Biol. 30:419-432. |
| 10. | Bird, D. F., and J. Kallf. 1984. Empirical relationships between bacterial abundance and chlorophyll concentrations in fresh and marine waters. Can. J. Fish. Aquat. Sci. 41:1015-1023. |
| 11. | Bjørnsen, P. K., B. Riemann, S. J. Horsted, T. G. Nielsen, and J. Pock-Sten. 1988. Trophic interactions between heterotrophic nanoflagellates and bacterioplankton in manipulated seawater enclosures. Limnol. Oceanogr. 33:409-420. |
| 12. | Cole, J., S. Findlay, and M. Pace. 1988. Bacterial production in fresh and salt water ecosystems: a cross system overview. Mar. Ecol. Prog. Ser. 43:1-10. |
| 13. |
De Rijk, P., and R. De Wachter.
1993.
DCSE, an interactive tool for sequence alignment and secondary structure research.
Comput. Appl. Biosci.
9:735-740 |
| 14. |
Fernandes, A. P.,
K. Nelson, and S. M. Beverley.
1993.
Evolution of nuclear ribosomal RNA in kinetoplastid protozoa: perspectives on the age and origin of parasitism.
Proc. Natl. Acad. Sci. USA
90:11608-11612 |
| 15. |
González, J. M.,
J. Iriberrri,
L. Egea, and I. Barcina.
1990.
Differential rates of digestion of bacteria by freshwater and marine phagotrophic protozoa.
Appl. Environ. Microbiol.
56:1851-1857 |
| 16. | Gunnison, D., and M. Alexander. 1974. Resistance and susceptibility of alga to decomposition by natural microbial communities. Limnol. Oceanogr. 20:64-70. |
| 17. | Hansen, L., G. F. Krog, and M. Søndergaard. 1986. Decomposition of lake phytoplankton. 1. Dynamics of short-term decomposition. Oikos 46:37-44. |
| 18. | Hasegawa, M., H. Kishino, and T. Yano. 1985. Dating of the human-ape splitting by a molecular clock of mitochondrial DNA. J. Mol. Evol. 4:406-425. |
| 19. | Jewel, W. J., and P. L. McCarty. 1971. Aerobic decomposition of algae. Environ. Sci. Technol. 5:1023-1031. |
| 20. | Kirchman, D. L., Y. Suzuki, C. Garside, and H. W. Ducklow. 1991. High turnover rates of dissolved organic carbon during a spring phytoplankton bloom. Nature 352:612-614. |
| 21. | Mudryk, Z., and W. Donderski. 1997. The occurrence of heterotrophic bacteria decomposing some macromolecular compounds in shallow estuarine lakes. Hydrobiologia 342/343:71-78. |
| 22. |
Muyzer, G.,
E. C. Dewaal, 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 |
| 23. |
Nei, M., and W. H. Li.
1979.
Mathematical model for studying genetic variation in terms of restriction endonucleases.
Proc. Natl. Acad. Sci. USA
76:5269-5273 |
| 24. | Nold, S. C., and G. Zwart. 1998. Patterns and governing forces in aquatic microbial communities. Aquat. Ecol. 32:17-35. |
| 25. |
Oró, J.,
T. G. Tornabene,
D. W. Nooner, and E. Gelpi.
1967.
Aliphatic hydrocarbons and fatty acids of some marine and freshwater microorganisms.
J. Bacteriol.
93:1811-1818 |
| 26. |
Ragan, M. A.,
C. L. Goggin,
R. J. Cawthorn,
L. Cerenius,
A. V. C. Jamieson,
S. M. Plourde,
T. G. Rand, and K. Söderhäll.
1996.
A novel clade of protistan parasites near the animal-fungal divergence.
Proc. Natl. Acad. Sci. USA
93:11907-11912 |
| 27. | Roessler, P. G. 1990. Environmental control of glycerolipid metabolism in microalgae: commercial implications and future research directions. J. Phycol. 26:393-399. |
| 28. | Sanders, R. W., D. A. Caron, and U.-G. Berninger. 1992. Relationships between bacteria and heterotrophic nanoplankton in marine and fresh waters: an inter-ecosystem comparison. Mar. Ecol. Prog. Ser. 86:1-14. |
| 29. |
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.
58:2381-2385 |
| 30. | Shifrin, N. S., and S. W. Chisholm. 1981. Phytoplankton lipids: interspecific differences and effects on nitrate, silicate and light-dark cycles. J. Phycol. 17:374-384. |
| 31. |
 imek, K., and T. H. Chraznowski.
1992.
Direct and indirect evidence of size-selective grazing on pelagic bacteria by freshwater nanoflagellates.
Appl. Environ. Microbiol.
58:3715-3720 |
| 32. |
Tranvik, L. J., and M. G. Höfle.
1987.
Bacterial growth in mixed cultures on dissolved organic carbon from humic and clear waters.
Appl. Environ. Microbiol.
53:482-488 |
| 33. | Upton, A. C., and D. B. Nedwell. 1989. Nutritional flexibility of oligotrophic and copiotrophic Antarctic bacteria with respect to organic substrates. FEMS Microbiol. Ecol. 62:1-6. |
| 34. |
van Hannen, E. J., and H. J. Gons.
1997.
UVC-induced lysis and detritus production of Oscillatoria limnetica in a two-stage continuous-flow system.
J. Plankton Res.
19:723-733.
|
| 35. | van Hannen, E. J., M. P. van Agterveld, H. J. Gons, and H. J. Laanbroek. 1998. Revealing eukaryotic genetic diversity in aquatic environments by denaturing gradient gel electrophoresis. J. Phycol. 34:206-213. |
| 36. | van Hannen, E. J., H. J. Gons, J. Ebert, H. L. Hoogveld, M. P. van Agterveld, G. Zwart, and H. J. Laanbroek. The microbial community structure in a large fresh water lagoon: an analysis using both flow cytometry and denaturing gradient gel electrophoresis. Submitted for publication. |
| 37. | van Hannen, E. J., M. Veninga, J. Bloem, H. J. Gons, and H. J. Laanbroek. Genetic changes in the bacterial community structure associated with protistan grazers. Arch. Hydrobiol., in press. |
| 38. |
van Hannen, E. J.,
G. Zwart,
M. P. van Agterveld,
H. J. Gons,
J. Ebert, and H. J. Laanbroek.
1999.
Changes in the bacterial and eukaryotic community structure after mass lysis of filamentous cyanobacteria associated with viruses.
Appl. Environ. Microbiol.
65:795-801 |
| 39. | Van Wambeke, F. 1994. Influence of phytoplankton lysis or grazing on bacterial metabolism and trophic relationships. Microb. Ecol. 27:143-158. |
| 40. | Verity, P. G. 1991. Feeding in planktonic protozoans: evidence for non-random acquisition of prey. J. Protozool. 38:69-76. |
| 41. | White, P. A., J. Kalff, J. B. Rasmussen, and J. M. Gasol. 1991. The effect of temperature and algal biomass on bacterial production and specific growth rate in freshwater and marine habitats. Microb. Ecol. 21:99-118. |
| 42. | Zwart, G., R. Huismans, M. P. van Agterveld, Y. Vande 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. |
Applied and Environmental Microbiology, June 1999, p. 2478-2484, Vol. 65, No. 6
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