We have used molecular biological methods to study the distribution
of microbial small-subunit rRNAs (SSU rRNAs), in relation to chemical
profiles, in offshore Lake Michigan sediments. The sampling site is at
a depth of 100 m, with temperatures of 2 to 4°C year-round. RNA
extracted from sediment was probed with radiolabeled oligonucleotides
targeting bacterial, archaeal, and eukaryotic SSU rRNAs, as well as
with a universal probe. The coverage of these probes in relation to the
present sequence database is discussed. Because ribosome production is
growth rate regulated, rRNA concentrations are an indicator of the
microbial populations active in situ. Over a 1-year period, changes in
sedimentary SSU rRNA concentrations followed seasonal changes in
surface water temperature and SSU rRNA concentration. Sedimentary depth
profiles of oxygen, reduced manganese and iron, and sulfate changed
relatively little from season to season, but the nitrate concentration
was approximately fivefold higher in April and June 1997 than at the
other times sampling was done. We propose that sediment microbial SSU
rRNA concentrations at our sampling site are influenced by seasonal inputs from the water column, particularly the settling of the spring
diatom bloom, and that the timing of this input may be modulated by
grazers, such that ammonia becomes available to sediment microbes
sooner than fresh organic carbon. Nitrate production from ammonia by
autotrophic nitrifying bacteria, combined with low activity of
heterotrophic denitrifying bacteria in the absence of readily
degradable organic carbon, could account for the cooccurrence of high
nitrate and low SSU rRNA concentrations.
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INTRODUCTION |
Molecular microbiological methods
based on nucleic acid (RNA and DNA) extraction can yield information
about the in situ distribution and activities of multiple microbial
groups simultaneously from a relatively small volume of sample. They
are especially useful for sediments, where numerous different species
are present and chemical composition and microbial populations can
change on a scale of millimeters to centimeters. Molecular methods are
also free of the well-known biases associated with traditional
culture-based approaches, such as the preferential recovery of species
that are well adapted to laboratory conditions, although issues such as
extraction efficiency have yet to be fully resolved.
In this study, we have used oligonucleotide probes to characterize
small-subunit rRNAs (SSU rRNAs) extracted from sediments at an offshore
Lake Michigan site. SSU rRNA is an essential component of the ribosome,
the RNA-protein complex responsible for protein synthesis in both
prokaryotes and eukaryotes. rRNA-encoding genes (rDNAs) from thousands
of species have now been sequenced and used to infer phylogenetic
relationships based on nucleotide sequence divergence
(67). Because the ribosome content generally increases with growth rate (reviewed in reference 33) and decreases
with starvation (see, e.g., references 14, 23, and
46), rRNA quantitation by oligonucleotide probe
hybridization can be used to estimate the composition of the actively
growing population at different levels of phylogenetic resolution.
Growth rate regulation of rRNA content differs among bacterial species
(5, 7, 16, 17, 21) and even strains (30),
however, so the precise relationship between SSU rRNA concentration and
community growth rate will depend on the species present
(32).
The activities of sediment microbes help determine how much of the
carbon and nitrogen that reaches the bottoms of lakes and other bodies
of water is buried and how much is returned to the water column or
atmosphere. Deepwater sediments, with a relatively constant dark and
cold physical environment, might be expected to have fairly constant
and low levels of microbial activity. However, evidence has been
accumulating that benthic sediments respond to seasonal changes nearer
the water surface (see, e.g., references 11, 49, and
62). We show here that microbial SSU rRNA concentrations
at our sampling site follow a seasonal pattern that appears to be
closely linked to seasonal changes in the water column. In particular,
we present evidence that nitrogen cycling in benthic Lake Michigan
sediments may have a strong seasonal component.
| |
MATERIALS AND METHODS |
Site description and sample collection.
The Fox Point
sampling site is approximately 27 km northeast of Milwaukee, Wis.
(latitude 43°11'40"N, longitude 87°40'11"W) at a depth of 
100 m.
The bottom water temperature is between 2 and 4°C year-round. The
sedimentation rate has been calculated as 0.24 cm
year
1 (19).
137Cs profiles suggest that the Fox Point
sediments are mixed to a depth of approximately 2 cm (50).
Sediment was collected with a 30-cm2 box corer
from the R/V Neeskay. Three-inch-diameter cylindrical subcores were taken for chemical analysis and nucleic acid extraction. The subcores were transported in an ice water bath or refrigerator protected from the light, stored in the dark at 4°C, and processed within 24 to 48 h. Prior to sectioning of a subcore for nucleic acid analysis, a disposable anaerobic glove bag (four-handed model; Sigma Chemical Co., St. Louis, Mo.) was taped over the end of the core
liner. The glove bag atmosphere was exchanged three times and replaced
with nitrogen, and slight positive pressure was maintained under
continuous flow. The core was pushed up out of the liner with a water
pressure-driven extruder and sectioned at 0.5-cm or greater intervals.
Core slices were transferred to glass screw-cap vials, kept on ice
under nitrogen until sectioning was completed, and then transferred to
an anaerobic hood for the initial steps of RNA extraction. Water
samples for RNA extraction were collected from depths of 0, 2, 5, 10, 15, 20, 25, 30, 60, 90, and 100 m with a Niskin bottle, transferred
to Cubitainers (Hedwin Corp., Baltimore, Md.), and processed on deck
within 1 to 12 h. Measured volumes of water (approximately 1 liter) were filtered through 0.2-µm-pore-size filters (Millipore
Corp., Bedford, Mass.) which were placed in 50-ml Falcon tubes and
frozen immediately on dry ice. Temperature, chlorophyll fluorescence,
and oxygen saturation were measured with a SeaBird instrument package.
Oxygen measurement.
Oxygen was measured in the sediment
cores with a Clark-style O2 microelectrode (model
737GC; Diamond General Corp., Ann Arbor, Mich.) positioned with a
micromanipulator and coupled to an analog chemical microsensor (Diamond General).
Chemical methods.
Pore water was recovered by squeezing
whole subcores into 5-ml plastic syringes (4). The water
was subsequently filtered through 0.2-µm Acrodisc HT Tuffyrn syringe
filters (Fisher Scientific). Nitrate and nitrite concentrations were
determined by flow injection analysis (automated cadmium reduction
method) (13). Sulfate was determined by ion chromatography
(13). Ammonium was measured using the indolphenol blue
method (34). All anion samples were run in duplicate
unless the sample volume was too low. Dissolved manganese(II) and
iron(II) were determined from pore water by atomic absorption
spectroscopy. Squeezer syringes were acidified with approximately 50 µl of 1 N HCl for preservation until chemical analysis according to
the manufacturer's instructions (IL Video 12 AAS; Allied Analytical
Systems, Andover, Mass.). Methane concentrations were measured by gas
chromatography using a flame ionization detector as described by Waples
(65).
RNA extraction and membrane hybridization.
For sediment
samples, 0.2-g aliquots of sediment were transferred with sterile 1-ml
disposable syringes to screw-cap Eppendorf tubes containing low-pH
buffer, buffer-equilibrated phenol (pH 5.1), sodium dodecyl sulfate,
and 0.5 g of zirconium beads (53), shaken for 1 min
with a Mini-Beadbeater (Biospec Products, Bartlesville, Okla.) in an
anaerobic hood, and then transported on ice to our laboratory. RNA was
isolated by bead beating, phenol-chloroform extraction, and ethanol
precipitation as previously described (42, 53). RNA was
resuspended in 200 µl of sterile deionized water. For water samples,
frozen filters were later crushed with sterile, baked spatulas and
transferred to screw-cap Eppendorf tubes and extracted as described for
sediment samples. RNA was transferred to nylon membranes in triplicate
and probed with radiolabeled oligonucleotides (Table
1) purchased from Operon Technologies Inc., Alameda, Calif. Membranes were prehybridized at 40°C and washed
at the temperatures listed. Hybridization was measured with a
PhosphorImager (model 400S or Storm; Molecular Dynamics Inc.,
Sunnyvale, Calif.).
DNA extraction and amplification and reverse transcription-PCR
(RT-PCR).
For the sequences designated LMBA, DNA was extracted
from 5-g sediment samples, collected in July and August 1993, by the method of Fuhrman et al. (22) and purified over a Chroma
Spin-100 column (Clontech Inc., Palo Alto, Calif.).
High-molecular-weight DNA was excised from a 0.6% agarose gel and
amplified using a 1605 Air Thermo-Cycler (Idaho Technology, Idaho
Falls, Idaho) with the Bacteria-specific primers Bact11F and
Bact1492AR (Table 2) according to the
manufacturer's instructions. DNA was denatured (94°C for 4 min) and
amplified for 30 cycles (92°C for 1 min, 50°C for 1 min, and 72°C
for 1 min 30 s), and products were extended (72°C for 5 min) to
facilitate cloning. Amplification products were cloned into the TA
vector (Invitrogen Corp., San Diego, Calif.). Isolated colonies were
streak purified and used to prepare plasmid DNA by an alkaline lysis
miniprep procedure (V. Schulz and R. Karls, personal communication).
Clones were designated LMBA (for Lake Michigan, bacterial
amplification, Alm) followed by a number.
The sequences designated LMBGA (for Bact11F-Geo825RA) were obtained
during an attempt to amplify Geobacter-like SSU rDNA
sequences as part of another study. DNA was isolated from sediment
collected on 3 August 1995 by lysozyme and freeze-thaw treatments,
phenol-chloroform extraction, and ethanol precipitation, essentially by
the method of Tsai and Olson (58). DNA was separated from
brown humic substances on a 1% agarose gel and purified from agarose
by centrifugation in 0.2-µm-pore-size SpinX centrifuge filter units
(Costar Inc., Cambridge, Mass.). Oligonucleotide primers were purchased
from Operon Technologies, Inc. DNA was amplified with a Hybaid PCR Express (Ashford, Middlesex, United Kingdom) according to the manufacturer's instructions, using the primer pair Bact11F-Geo825RA (Table 2). DNA was denatured (94°C for 30 s) and amplified for 30 cycles (92°C for 15 s, 53.3°C [calculated temperature]
for 15 s, and 72°C for 45 s), and products were extended
(72°C for 1 min) to facilitate cloning. Amplification products were
cloned into the pCR4-TOPO vector (Invitrogen). Isolated colonies were streak purified and used to prepare plasmid DNA. Clones designated LMRTGA were obtained by RT-PCR of sediment-extracted RNA, collected on
3 August 1995, with the Access RT-PCR kit (Promega Inc., Madison, Wis.)
using the primer pair Bact11F-Geo825RA. PCR was performed as described
for the LMBA clones, and products were cloned in the pCR4-TOPO cloning
vector for sequencing.
An automated DNA sequencer (model 4000L; LiCor Corp., Lincoln, Nebr.),
a SequiTherm Long-Read Kit (LC) (Epicentre Technologies Inc.,
Madison, Wis.), and M13Fwd(-29) and M13Reverse IRD41-labeled primers (LiCor) were used for DNA sequencing. Sequence analysis was
done using the Arb sequence database analysis package
(55).
Nucleotide sequence accession numbers.
The SSU rRNA
sequences presented here have been deposited in GenBank under accession
numbers AF320917, AF320919 to -23, AF320930, AF320955 and -56, and
AF320972 to -75.
| |
RESULTS |
Chemical profiles and universal probe hybridization.
Sediment chemistry profiles of the Fox Point sediment cores generally
showed the expected progression attributed to microbial utilization of
terminal electron acceptors in order of their energy yield: oxygen
depletion, nitrate depletion, Mn(II) accumulation, Fe(II) accumulation,
and sulfate depletion (Fig. 1). The
concentrations and depth profiles of oxygen, sulfate, Mn(II), Fe(II),
and ammonia were relatively constant from sample to sample, where
measured. Nitrate concentrations, however, were severalfold higher in
April and June 1997. Sediment surface SSU rRNA concentrations also
varied, ranging from 53 ng g of sediment1 in
April 1997 to 1,745 ng g of sediment1 in
September 1996. Thus, nitrate concentrations were highest when rRNA
concentrations were lowest.
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FIG. 1.
Sediment chemistry and universal probe
(S-*-Univ-1390-a-A-18) hybridization. Chemical and SSU rRNA
concentrations were determined as described in Results.  , 2 May
1996;  , 4 September 1996;  , 13 November 1996;  , 2 April 1997;
 , 5 June 1997. Nitrite and oxygen data for April and ammonia data
for June are not available.
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Considering all samples for which both nitrate and rRNA concentrations
were measured, nitrate concentrations of greater than or equal to 50 µM were found only with SSU rRNA concentrations of less than 100 ng g
of sediment1 (Fig.
2). There was a positive correlation
between nitrate and SSU rRNA concentrations for the
May, September, and November samples (correlation coefficient = 0.607; P < 0.01) (52), which is
unsurprising since both of these parameters generally decrease with
depth, but there was no significant correlation for the April and June samples or for all samples taken together.
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FIG. 2.
Universal probe hybridization and nitrate
concentration. , 2 May 1996; , 4 September 1996; , 13 November
1996; , 2 April 1997; , 5 June 1997. There is a positive
correlation between nitrate concentration and SSU rRNA concentration
for the May, September, and November samples (correlation
coefficient = 0.607; P < 0.01)
(52) but no significant relationship for the April and
June samples.
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The nitrate distribution in June was unusual, being very low at the
sediment surface and peaking at a depth of around 2 cm (Fig. 1). The
nitrite concentration was also relatively high compared to that in
September and November (nitrite was not measured in April). The SSU
rRNA concentration in the 0- to 0.5-cm interval of this June sample was
414 ng g of sediment1, compared to 53 ng g of
sediment1 in April, while the concentrations at
0.5 to 1 cm were 34 and 55 ng g of sediment1,
respectively. We have some evidence that the June profile may have been
caused by some localized event (see Fig. 7; discussed below). One
possibility is that a recent nutrient input resulted in increased
near-surface microbial respiration and nitrogen demand, reflected in
rRNA concentration, which consumed the near-surface nitrate.
Comparison of sediment and water column rRNA concentrations.
Sedimentary SSU rRNA concentrations appeared to be related to seasonal
changes in the overlying water column (Fig.
3). They were highest in the stratified
season (September) and lowest in early spring (April). Where water
column RNA data are available, seasonal trends are also apparent, with
highest concentrations in late fall (November) and lowest
concentrations in early spring (April). Water column SSU rRNA
concentrations for July and October 1997 were higher and more
stratified by depth than those for the sampling times reported here
(43, 63), in accordance with other seasonal cycles in the
lake. Surface water temperatures ranged from 22.0°C in September to
1.8°C in April, while bottom water temperatures varied only between
1.8 and 4.0°C. Sediment SSU rRNA concentrations were highest in
September, when water rather than sediment temperatures were highest.
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FIG. 3.
Comparison of water column (A) and sediment (B) SSU rRNA
concentrations. RNA extracted from water or sediment samples was
quantitated by membrane hybridization with the universal probe
(S-*-Univ-1390-a-A-18). Water temperature was measured with the SeaBird
instrument package.
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Domain-level probe hybridization.
Hybridization to all three
domain-level probes (bacterial, eukaryotic, and archaeal), as well as
to the universal probe, was lowest in April 1997 and highest at most
depths in either September or November 1996 (Fig.
4). Note that different horizontal scales are used for each of the probes. Bacterial SSU rRNA predominated, except in April, followed by eukaryotic rRNA. Archaeal rRNA was a small
fraction of the total at all times and depths sampled. For a given
sample, the depth distributions for all four probes were similar.
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FIG. 4.
Domain-level probe hybridization. RNA extracted from
each 0.5- or 1-cm depth interval was transferred to nylon membranes and
quantitated with a radiolabeled universal (S-*-Univ-1390-a-A-18),
bacterial (S-D-Bact-0338-a-A-18), eukaryotic (S-D-Euca-1379-a-A-16), or
archaeal (S-D-Arch-0915-a-A-20) oligonucleotide probe. Note that a
different horizontal scale was used for each probe.
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Hybridization to the archaeal probe was generally greatest in the oxic
to suboxic zone. This is contrary to our initial expectation that
archaeal rRNA would be contributed primarily by obligately anaerobic
methanogens in the anoxic deeper sediments. Some of this archaeal
signal can likely be attributed to the cold-water crenarchaeota
(42; B. J. MacGregor, unpublished observations), which have been found in other oxic habitats (see, e.g., reference 12). Hybridization to probes targeting several different
groups of methanogenic archaea has also been found in Fox Point samples from the oxic zone, suggesting the possibility of anaerobic niches there (1).
Ideally, hybridization to the universal probe should equal the sum of
bacterial, eukaryotic, and archaeal probe hybridizations. Domain
summations for May, September, and November were generally between 40 and 120% (Fig. 5). Both the lowest and
the highest summations were obtained with low-rRNA samples collected
from deep in the sediment or in April and June, and they may be due in
part to uncertainties in measuring very low RNA concentrations. Chloroplast rRNA might also contribute to low domain summations (discussed below). Domain summations of greater than 100% may result
from RNA degradation: the universal probe target site is located in a
region of the SSU rRNA molecule with relatively little secondary
structure (68) and may be especially subject to
degradation (48). While RNA degradation is generally
attributed to laboratory RNase contamination, nucleases are of course
also found in natural environments. Partially degraded RNA might be
especially abundant where cell lysis is frequent, due, for example, to
grazing.
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FIG. 5.
Domain summations. Hybridization to the bacterial,
eukaryotic, and archaeal probes is expressed as a percentage of
hybridization to the universal probe. Two summations over 200% are
shown in the inset graphs. Gaps indicate depth intervals for which RNA
extractions were not done.
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Bacterial SSU rRNA predominated, with several exceptions. The two
deepest samples from May 1996 and the 15- to 16-cm sample from
September hybridized only to the archaeal probe, suggesting that
archaeal methanogens may be active at these greater depths. In April,
eukaryotic rRNA was most abundant, although as a percentage of
universal probe hybridization it was not more abundant than in other
months. Eukaryotic rRNA was also proportionately abundant in December
1997 (see Fig. 7C) and between depths of 0.5 and 1.5 cm in box core 1 from June 1997 (Fig. 5; see Fig. 7B). The nitrate concentration and
eukaryotic probe hybridization as proportions of total domain-level SSU
rRNA were positively correlated overall (Fig.
6) (correlation coefficient, 0.402;
P < 0.01) (52). Samples in which only
eukaryotic RNA was detected (all of which were collected in April and
June 1997) had a wide range of nitrate concentrations, however. An
explanation for the cooccurrence of high nitrate levels and
predominance of eukaryotic rRNA is suggested in Discussion.
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FIG. 6.
Nitrate concentration and proportion of eukaryotic SSU
rRNA. The eukaryotic rRNA concentration is expressed as a proportion of
the combined eukaryotic, bacterial, and archaeal probe hybridization.
, 2 May 1996; , 4 September 1996; , 13 November 1996; , 2 April 1997; , 5 June 1997.
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Spatial variability in probe hybridization.
Duplicate box
cores and subcores were analyzed on three occasions to explore the
degree of spatial as opposed to seasonal variation in the sediment
community. RNA recovery from the 2 April 1997 box cores was low, and
absolute site-to-site differences were small (Fig.
7A [note that horizontal scales differ
from sampling time to sampling time and for the
archaeal probe as opposed to the others]). Nearly all of the rRNA
detected was eukaryotic. Replicate cores taken on 5 June 1997 differed
(Fig. 7B). In box core 1, eukaryotic SSU rRNA accounted for nearly all
probe hybridization in the 0.5- to 1.5-cm interval. Eukaryotic probe
hybridization in box core 2 was more evenly distributed as a proportion
of universal probe hybridization, but it was also greatest between 0.5 and 1.5 cm. This suggests a eukaryotic population (perhaps seasonal) concentrated at this depth. Subcore 1A was used for the full-length June hybridization profiles discussed above, and box core 1 was used
for the chemical measurements, which showed high nitrate concentrations. The higher proportion of bacterial rRNA found in the
summer and fall samples may develop unevenly across the lake floor,
perhaps controlled by local rates of algal delivery and grazing.
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FIG. 7.
Replicate box cores. Two box cores were collected
from sites approximately 1 km apart, and cylindrical subcores were
taken for RNA extraction from the top 2.5 cm of sediment. Extracted RNA
was hybridized with the universal and domain probes. Domain-level
results are displayed both as concentrations and as percentages of
universal probe hybridization. Because RNA concentrations ranged from
picograms to micrograms per gram of sediment, different horizontal-axis
scales were used for each sampling time and for the archaeal versus the
universal, bacterial, and eukaryotic probes.
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The depth profiles of probe hybridization in the 17 December 1997 samples (Fig. 7C) were generally similar to each other, except that
universal probe hybridization in the 1- to 2-cm interval of subcore 1A
was severalfold higher than that in the other samples. Subcores 1A and
1B were taken from adjacent positions in box core 1, so this represents
variability on a scale of <20 cm. Most of the difference is
attributable to increased eukaryotic probe hybridization. Our sample
size of one to several grams is not sufficient for representative
sampling of larger eukaryotes, so peaks such as this are not
unexpected. There was a peak of archaeal probe hybridization at the
same depth, suggesting the possibility of a symbiotic association such
as those found in anaerobic ciliates (18).
December oxygen profiles (Fig. 8) were
measured on subcores separate from those used for RNA extraction. In
box core 1, a subcore recovered from closer to subcore 1A than to
subcore 1B was used. Oxygen penetration was approximately 1 cm deeper
in core 1 than in core 2 or in any other of the Fox Point cores sampled to date (Fig. 1 and unpublished observations). The combination of a
high proportion of eukaryotic probe hybridization and increased oxygen
penetration suggests bioturbation and perhaps a response by microscopic
eukaryotes to increased oxygen availability.
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FIG. 8.
Replicate oxygen profiles from samples taken on 17 December 1997. Oxygen concentrations were measured by microelectrode on
subcores taken from each of the two box cores.
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Detection of potential nitrifying bacteria.
The most likely
source of high sediment nitrate concentrations is biological oxidation
of ammonia. Although we have not yet quantified known nitrifier groups,
we identified a new clade of organisms closely affiliated with known
beta-proteobacterial nitrifiers (Fig. 9).
These sequences were recovered from several different sediment samples
using two different primer sets not specifically targeting nitrifiers,
suggesting a high abundance of this novel clade. A closely related
sequence, MNH4, was isolated from Green Bay (Lake Michigan) sediments
(Fig. 9) (54).
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FIG. 9.
Beta-proteobacterial SSU rRNA sequences amplified from
Lake Michigan sediment. Sequences in boldface with two asterisks were
recovered from Fox Point sediments in this study. Sediment sampling
intervals are shown in square brackets. Single asterisks indicate
sequences amplified from Green Bay sediment by Stein et al.
(61). Numbers in parentheses indicate the number of
sequences included in a particular branch. The tree was calculated
using the neighbor-joining method, as implemented in the sequence
analysis program Arb (55). Gram-positive bacterial SSU
rRNA sequences were used to root the tree. The scale bar represents one
fixed mutation per 10 nucleotide positions.
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DISCUSSION |
Probe coverage.
Because the probes used were designed several
years ago, and the number of SSU rRNA sequences available from GenBank
is increasing rapidly, we first compare the probes to the present
sequence database and show that their coverage remains substantially complete.
(i) Universal and bacterial probes.
Bacterial rRNA accounted
for the majority of probe hybridization in the Fox Point samples, so
differences in probe coverage between the universal
(S-*-Univ-1390-a-A-18) and bacterial (S-D-Bact-0338-a-A-18) probes are
a particular concern. Most bacterial groups presently recognized
contain only a small proportion of nontarget species for either probe
(Fig. 10 [due to
computer limitations, only species missed by one or
both probes are included]). Some of the mismatches, particularly the
phylogenetically isolated ones, may be due to sequencing errors. The
greatest concentration of universal probe mismatches is found in the
epsilon proteobacteria, while bacterial probe mismatches are especially
prevalent among the chlamydiales and planctomycetales (as a proportion
of the presently described species). Outside of the deeply branching
Thermotogales and Aquificales, few bacterial
species are missed by both probes. The universal probe mismatches
should be partially compensated for by the wash temperature (Table 1),
which was experimentally chosen to allow detection of single-mismatch
archaeal species without overrepresentation of the Bacteria
and Eukarya (69). Daims et al.
(10) have recently designed a set of probes that together
target all presently known bacterial SSU rRNA sequences.
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FIG. 10.
Bacterial species with mismatches to the
universal (S-*-Univ-1390-a-A-18) and/or bacterial
(S-D-Bact-0338-a-A-18) probe. The universal and bacterial probe
sequences were checked against a collection of over 5,700 nearly
full-length aligned prokaryotic SSU rRNA sequences, which include
target regions with no ambiguities for both probes. Only species with
mismatches to one or both probes are included in the tree shown here,
which was calculated by neighbor joining with Olsen distance correction
using the Arb sequence database program (55). Archaeal SSU
rRNA sequences were used to root the tree. The scale bar represents one
fixed mutation per 10 nucleotide positions.
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(ii) Eukaryotic probe.
Eukaryotic rRNA was also a large
proportion of the total in most of the Fox Point samples. Of 1,946 nearly full-length eukaryotic SSU rDNA sequences obtained from GenBank,
92 have mismatches to the eukaryotic probe (S-D-Euca-1379-a-A-16) only,
65 have mismatches to the universal probe only, and 19 have mismatches
to both probes. Of aquatic microeukaryotes, rDNA sequences from the
Euglenales in particular match the universal but not the
eukaryotic probe. Without more information about the microeukaryotic
populations at Fox Point, we cannot evaluate the possible contribution
of these species to the domain summations.
(iii) Archaeal probe.
Nearly all archaeal SSU rDNA sequences
have a perfect target site for the archaeal probe
(S-D-Arch-0915-a-A-20) and a single mismatch with the universal probe,
which is compensated for by a decreased wash temperature
(69). One cluster of species with a single mismatch to the
archaeal probe is found among the thermophiles (Fig.
11); the five other exceptional species
are scattered among the archaeal clades. Archaeal probe hybridization
generally represents only a small proportion of the total, so it should
be a negligible source of error in any case.
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FIG. 11.
Archaeal probe coverage. All of the sequences shown are
targeted by both probes unless otherwise indicated. The tree was
calculated by neighbor joining with Olsen distance correction using the
Arb sequence database program (55). Bacterial SSU rDNA
sequences were used to root the tree. The scale bar represents one
fixed mutation per 10 nucleotide positions.
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(iv) Organellar rRNA.
Mitochondria and chloroplasts, which are
believed to have evolved from endosymbiotic bacteria, contain
rRNA-encoding genes. Although the Fox Point sampling site is below the
photic zone, the chloroplasts of photosynthetic organisms that settle
to the bottom may contribute to the rRNA pool. Of 70 nearly full-length algal and higher-plant chloroplast rRNA sequences examined (Fig. 12), 39 are targeted by both the
universal and bacterial probes, 24 are targeted by the universal probe
only, 6 are targeted by the bacterial probe only, and 1 is targeted by
neither probe. Chloroplast rRNAs that hybridized only with the
universal probe might help account for some of the low domain
summations, particularly in April 1997 (Fig. 5).
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FIG. 12.
Bacterial and universal probe coverage of chloroplast
SSU rDNA sequences. All sequences shown are targeted by both probes,
unless otherwise indicated. The tree was calculated by neighbor joining
using the Arb sequence database program (55). A
delta-proteobacterial SSU rRNA sequence was used to root the tree. The
scale bar represents one fixed mutation per 10 nucleotide positions.
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Of 184 mitochondrial SSU rDNA sequences checked, nearly all had
multiple mismatches to the universal probe and all of the domain probes
(data not shown) and thus should have little impact on the domain
summations. The only two sequences with perfect matches to any probe
are both from aquatic species, however. The mitochondrial rDNAs of the
red alga Porphyra purpurea and the freshwater amoeba
Acanthamoeba castellani Neff include universal probe target sites.
Seasonal changes in SSU rRNA and nitrate concentration: a suggested
role for grazers.
Because the temperature of the Fox Point
sediments is nearly constant year-round, as were previously measured
chemical profiles (K. H. Nealson, unpublished observations), we
expected that microbial SSU rRNA concentrations would show little
evidence of seasonality. However, we found at least a 10-fold variation
in the upper sediment layers during the course of a year. We also found
that while sulfate, iron, manganese, and oxygen profiles changed
little, nitrate concentrations were severalfold higher in spring.
Bottom water nitrate concentrations in southern Lake Michigan are
between 5 and 30 µM year-round (8) and are lower in
spring than in summer (Fig. 13), so
this difference is unlikely to be caused by direct nitrate input from
the water column. High nitrate levels were associated with low
concentrations of SSU rRNA (Fig. 2).
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|
FIG. 13.
Bottom water nitrite plus nitrate concentrations (data
are from reference 59). Station 1115 GLSB/L. MICH 17 was
chosen as the long-term Lake Michigan study site most similar to our
Fox Point station. It is located offshore of Racine, Wis., at
42°44'0"N, 87°25'0"W; the water depth is 350 ft (107 m). Only
measurements made at depths of greater than 80 m are shown. ,
1988; , 1989; , 1990;  , 1991;  , 1992; , 1993;  , 1996;
 , 1997.
|
|
Nutrient availability is probably the primary seasonal factor at the
Fox Point site. Sediment microbial communities can respond quickly to
the settling of real or simulated algal blooms (reviewed in reference
26). Cold, unstratified water and seasonally low grazer
populations allow a large proportion of the Lake Michigan spring diatom
bloom to settle directly to the sediment (24, 51). Brooks
and Torke (9) measured chlorophyll at Fox Point in 1973 and 1974. Integrated over the whole water column, the chlorophyll
concentration began to increase in late winter, increased more quickly
in March, and reached maximums in late May or early June each year.
Those authors proposed that silica depletion and/or the onset of
stratification may end the bloom in different years. Late-season
primary production generally reaches the bottom of the lake only
slowly. Nonsiliceous species sink more slowly than diatoms, and thermal
stratification forms a density barrier. Thus, diatoms dominated the
phytoplankton in deep sediment traps at a 100-m site in southeastern
Lake Michigan even in summer, when they were only a small proportion of
the algal population (51). Other seasonally varying
sediment inputs may include thermal bar transport (15, 40,
44), calcium carbonate precipitation (whiting) events (28,
57, 61), vertical migrations by animals, lateral transport by
deep currents, and anthropogenic inputs.
What might account for a combination of high nitrate concentration and
low microbial SSU rRNA concentration in the spring? We suggest that
benthic eukaryotes may modulate the timing of carbon and nitrogen
delivery to the sediment microbial community. Eukaryotic grazers are
proposed to quickly consume the bulk of the easily degraded new carbon
and to excrete ammonia. The ammonia can be converted to nitrate by
autotrophic nitrifying bacteria. Because of their generally low growth
yield, a small number of nitrifiers can achieve a high conversion rate
without causing a large increase in sediment SSU rRNA concentration.
Nitrate consumption by heterotrophic denitrifiers may then await the
slow hydrolysis of more recalcitrant carbon compounds. Preliminary
modeling results (not shown) suggest that this mechanism is
qualitatively possible. It may account for the seasonal association of
high nitrate concentrations with a high proportion of eukaryotic SSU
rRNA (Fig. 6), which could be contributed by diatoms (to the extent
that they retain their rRNA) and the animals grazing on them.
One prediction from this scenario is that oxygen might be more quickly
depleted when nitrifiers are active, because oxygen is required for
ammonia oxidation. Oxygen profiles were unfortunately not obtained in
April 1997, due to electrode malfunction. Oxygen concentrations
measured on 22 April 1995 (not shown) decreased from 100% saturation
at the sediment surface to 7% at the 1-cm depth, the steepest gradient
yet found at Fox Point, but as there was no further decrease from 1 to
3 cm, this measurement may not be reliable.
Studies on soil, sediment, and aquatic systems have found both
increased (29, 47, 56, 64) and decreased (37,
38) nitrification rates and/or nitrate concentrations in the
presence of eukaryotes. Because the source environments, animals
present, and laboratory conditions varied, it is difficult to draw
general conclusions. Roles proposed for animals include mixing of
ammonia into the oxic zone (64), ammonia production
(47), and selection for bacterial aggregates with lower
nitrification rates (37).
Benthic eukaryotes in Lake Michigan include Diporeia
(Pontoporeia) hoyi (35, 45),
Mysis relicta (opossum shrimp) (39), chironomid
(midgefly) larvae (66), and Stylodrilus
heringianus (36). The amphipod D. hoyi is
the dominant benthic invertebrate (35, 45). We observed
D. hoyi burrowing rapidly to and from a depth of several
centimeters in most Fox Point sediment cores. D. hoyi is a
detritivore, feeding in the top 2 cm of sediment, and may consume a
major share of fresh detritus (20, 24). Gardner et al.
(25) incubated gamma-irradiated or untreated Lake Michigan
sediments with or without added D. hoyi. Ammonia accumulated
significantly only in the treatment with animals and sterilized
sediment, implying that ammonia was excreted by D. hoyi and
then used by sediment organisms. Nitrate accumulated in the treatment
with animals and unsterilized sediment, suggesting nitrifier activity,
but the accumulation was small relative to the amount of ammonia
production inferred from the other treatments. Flux measurements on
separate cores showed that denitrification was significant and so
probably accounted for the remainder of the nitrogen. Repeating these
experiments with Fox Point sediments collected at different times of
the year, or with and without added fresh detritus, might indicate
whether grazers can in fact modulate the rate of nitrogen versus carbon input.
Conclusion.
Microbial rRNA concentrations in the sediment at
the 100-m-deep Fox Point sampling site appear to depend on seasonal
production nearer the surface. Although most physical and chemical
profiles measured were relatively constant over the 1-year sampling
period, SSU rRNA concentrations increased more than 10-fold between
early spring and fall. Community composition changed as well, with
eukaryotic rRNA predominating in winter (December) (Fig. 7C) and early
spring (April) (Fig. 7A) and bacterial rRNA predominating the rest of the year.
The cooccurrence of high nitrate and low SSU rRNA concentrations in
early spring was an unexpected finding. We propose that eukaryotes
grazing on settled diatoms may release spring-bloom nitrogen, as
ammonia, while consuming most of the readily degradable carbon
compounds. Nitrate produced from this ammonia by slow-growing autotrophic nitrifying bacteria may then accumulate until enzymatic hydrolysis makes more recalcitrant carbon sources available to heterotrophic denitrifiers.
Further experiments are needed to test this model. An attempt should be
made to isolate members of the beta-proteobacterial clade identified by
16S rDNA and rRNA amplification (Fig. 9) to determine whether they are
nitrifiers, as the most closely related species are. Oligonucleotide
probes and primers targeting the 16S rRNAs (reviewed in reference
60) and ammonia oxidation genes (reviewed in reference
6) of known nitrifying bacteria, as well as this new
group, could then be used to look for seasonal trends in nitrifier
activity. These could be related to surface production deposition by
sediment trap studies or by measuring chlorophyll and pigment
concentrations in sediment samples (62).
We thank Captain Ron Smith, First Mate Greg Stamatelakys, and all
of the crew of the R/V Neeskay for many pleasant (and
some exciting) cruises. Michael Leonardo helped with sediment sampling. Benjamin Van Mooy, Changrui Gong, Joseph Werne, and David Hollander helped to collect water samples. Michael Dollhopf made most of the
oxygen measurements. Thanks go to Art Brooks and the reviewers for helpful comments on the manuscript. The Storm PhosphorImager is
located in the Keck Biophysics Facility, Northwestern University.
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Abstract
Introduction
Present address: Pacific Northwest National Laboratories, Richland,
WA 99532.
Present address: Department of Geology and Geophysics, University
of Wisconsin, Madison, WI 53706.
Present address: Jet Propulsion Laboratory, California Institute
of Technology, Pasadena, CA 91109.
