Download Cell Size Distributions of Soil Bacterial and Archaeal

Cell Size Distributions of Soil Bacterial and Archaeal Taxa
Maria C. Portillo,a Jonathan W. Leff,a,b Christian L. Lauber,a Noah Fierera,b
Cooperative Institute for Research in the Environmental Sciences, University of Colorado, Boulder, Colorado, USAa; Department of Ecology and Evolutionary Biology,
University of Colorado, Boulder, Colorado, USAb
Cell size is a key ecological trait of soil microorganisms that determines a wide range of life history attributes, including the efficiency of nutrient acquisition. However, because of the methodological issues associated with determining cell sizes in situ, we
have a limited understanding of how cell abundances vary across cell size fractions and whether certain microbial taxa have consistently smaller cells than other taxa. In this study, we extracted cells from three distinct soils and fractionated them into seven
size ranges (5 ␮m to 0.2 ␮m) by filtration. Cell abundances in each size fraction were determined by direct microscopy, with the
taxonomic composition of each size fraction determined by high-throughput sequencing of the 16S rRNA gene. Most of the cells
were smaller than cells typically grown in culture, with 59 to 67% of cells <1.2 ␮m in diameter. Furthermore, each size fraction
harbored distinct bacterial and archaeal communities in each of the three soils, and many of the taxa exhibited distinct size distribution patterns, with the smaller size fractions having higher relative abundances of taxa that are rare or poorly characterized
(including Acidobacteria, Gemmatimonadetes, Crenarchaeota, Verrucomicrobia, and Elusimicrobia). In general, there was a direct relationship between average cell size and culturability, with those soil taxa that are poorly represented in culture collections
tending to be smaller. Size fractionation not only provides important insight into the life history strategies of soil microbial taxa
but also is a useful tool to enable more focused investigations into those taxa that remain poorly characterized.
T
he soil environment harbors an amazing diversity of microorganisms, with thousands of bacterial and archaeal taxa living
in individual soil samples (1). These taxa not only have a broad
array of physiologies and life history strategies but also can exhibit
a wide range of morphologies. In particular, we know that cell size
can vary considerably across bacterial and archaeal taxa, from 0.2
to 750 ␮m in diameter (2). This variation in cell size has important
effects on the physiological attributes of cells and their interactions with the soil environment, as cell diameter is inversely related to the surface area-to-volume ratio of cells. The surface areato-volume ratio is a key determinant of a wide range of cell
attributes, including the rate at which cells can take up nutrients
from their environment, maintenance energy requirements,
growth rates, and rates at which cells can release waste products
(3). Despite cell size being a key characteristic of soil microorganisms, we have a limited understanding of microbial size distributions in soil and how cell size may vary across the wide diversity of
taxa commonly found in soil.
We know from culture-based studies that individual microbial
taxa can exhibit a range of cell sizes depending on environmental
conditions and growth stage, with more actively growing cells living in resource-rich environments typically having larger cell diameters (4, 5). Nutrient deprivation has been shown to reduce cell
size (3), and not surprisingly, cell sizes in soil are generally considered to be smaller than cell sizes typically measured for the most
commonly studied isolates grown under laboratory conditions (4,
6). For example, previous work has shown that the majority of soil
bacteria are less than 0.5 ␮m in diameter (6), with some bacterial
cells small enough to pass through 0.2-␮m filters (7). These
“dwarf” cells appear to be very difficult to cultivate in the laboratory (8) and tend to grow more slowly than larger bacterial taxa
(9). There is also some evidence that specific bacterial and archaeal
taxa may be relatively more abundant in the smaller cell size fractions (10, 11), with many of these taxa likely retaining their small
size after cultivation (6, 10, 12). However, what remains to be
determined is if the unique cell size fractions generally represent
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Applied and Environmental Microbiology
distinct communities. In other words, we do not know how the
intertaxon variability in cell size compares to the cell size variability observed within individual taxa. We hypothesize that cell size
has a strong taxonomic signal, with certain taxa having smaller
cells than other taxa, a pattern that could result from either consistent taxonomic differences in cell growth rates or genetic constraints on cell size that contribute to differences in cell sizes regardless of growth conditions.
The knowledge gaps in our understanding of cell size distributions in soil persist because it is not easy to quantify variation in
cell size in environmental samples. Although cell size can readily
be determined for cultured isolates, the vast majority of soil microbial taxa are difficult to cultivate (13), and even for those taxa
that can be cultured, cell size distributions in culture may have
little to no bearing on the size distributions in soil due to differences in resource availability and environmental conditions (3–5).
Although sequence-based approaches have revolutionized our
ability to describe microbial diversity and the physiological attributes of individual taxa, such methods are of limited utility for
determining cell sizes. Likewise, although microscopy-based approaches, including fluorescent in situ hybridization, can be used
to determine the morphologies of soil microbial taxa that are resistant to culturing (e.g., see reference 14), the logistical difficulties
associated with these approaches make it difficult to gain detailed
information on how the wide range of taxa found in soil vary with
respect to their cell size distributions. To circumvent these limita-
Received 13 August 2013 Accepted 23 September 2013
Published ahead of print 27 September 2013
Address correspondence to Noah Fierer, [email protected].
Supplemental material for this article may be found at http://dx.doi.org/10.1128
/AEM.02710-13.
Copyright © 2013, American Society for Microbiology. All Rights Reserved.
doi:10.1128/AEM.02710-13
p. 7610 –7617
December 2013 Volume 79 Number 24
Size Distributions of Soil Bacterial and Archaeal Taxa
TABLE 1 General description of the collection sites, edaphic characteristics, and cell number counts from the three soil types studieda
Site
Latitude
(oN)
Longitude
(oW)
Prairie
39.1
96.6
Forest
Agricultural
41.9
42.4
72.1
85.4
a
Dominant plant species
pH
%C
%N
% silt ⫹
clay
Soil moisture
(g H2O g dry soil⫺1)
No. of cells
g dry soil⫺1
Andropogon gerardii, Schizachyrium
scoparium
Quercus rubra, Acer rubrum
Solanum tuberosum
6.84
3.74
0.30
78
0.31
1.8 ⫻ 1010
3.91
5.15
4.87
0.60
0.32
0.06
47
26
0.62
0.26
9.4 ⫻ 1010
2.1 ⫻ 1010
Cell counts were obtained on whole soils without any fractionation of cells by size. Soil texture was determined as described previously (1).
tions, we developed an approach for this study that involves isolating the microbial cells from a soil matrix using a Nycodenz
density gradient and then passing the cells through a series of
filters with pore sizes ranging from 5 to 0.2 ␮m. Cell abundances
on each of the filters were then determined via microscopy, with
the taxonomic identities of the cells in each size fraction determined by high-throughput sequencing of the 16S rRNA gene.
Using this combined approach, we were able to determine the size
distributions in total cell abundances across the soils and the size
distributions of individual bacterial and archaeal taxa to determine if some taxa are consistently larger or smaller than other
taxa.
MATERIALS AND METHODS
Soil sample collection and processing. Surface mineral soils (0- to 5-cm
depth, no O-horizon material) were collected from a native grassland
(tallgrass prairie) in eastern Kansas, a humid deciduous forest in Connecticut, and a cultivated agricultural field located at the Kellogg Biological
Station in Michigan. These three soils (which we refer to as prairie, forest,
and agricultural soils, respectively) were chosen because they represent a
range of soil types and site characteristics (Table 1). Our objective was not
to compare how cell size distributions vary across gradients in soil or site
characteristics but rather to determine the size distributions of microbial
taxa within each of these three distinct soils. All soils were sieved to 2 mm,
homogenized, and stored at 4°C until further processing.
Cell separation and pore size fractionation. Four 5-g replicate subsamples of each soil were placed into 50-ml sterile conical tubes, and 5 ml
of a cell fixing solution (50 mM tetrasodium pyrophosphate [TSP; pH
8.0], 4% formaldehyde, and 0.5% Tween 80, as per reference 15) was
added to each tube. The resultant slurries were incubated overnight at 4°C
to ensure that cells were inactivated to prevent changes in the microbial
communities during the cell separation and fractionation procedures. Soil
slurries were then vortexed for 15 min, with the larger soil particles allowed to settle to the bottom of the conical tube. Next, 1-ml aliquots of the
slurry were aseptically transferred to sterile 2-ml Eppendorf tubes containing 0.5 ml of Nycodenz (Axell, Westbury, NY) solution (80%, wt/vol,
prepared in 50 mM sterile TSP buffer), with the Nycodenz density gradient used to isolate bacteria from soil particles by following a procedure
described previously (16). Tubes were centrifuged at 10,000 ⫻ g for 90
min, after which the upper and middle cell-containing phases were transferred to new sterile 2-ml tubes with 1 ml of the 50 mM TSP buffer. After
mixing, the tubes were centrifuged at 20,000 ⫻ g for 15 min. The supernatant was discarded and the cell pellet was resuspended in 0.6 ml of the 50
mM TSP buffer containing 4% formaldehyde. The volume of suspended
cells from each sample replicate was pooled, and approximately 5 ml of
cell suspension was obtained for each of the four replicates per sample. Six
negative-control samples (consisting of 50 mM TSP buffer instead of soil
slurry) were processed through the Nycodenz density gradient described
above to check for any contamination.
Fractionation of microbial cells based on size was accomplished by
passing the cell suspension through polycarbonate membrane filters with
successively smaller pores (5, 3, 1.2, 0.8, 0.6, 0.4, and 0.2 ␮m) using a
gentle vacuum (⬍20,000 Pa). The filters as well as 0.5 ml of the flow-
December 2013 Volume 79 Number 24
through from each filter were retained and stored at ⫺20°C for community analysis. An additional 0.1 ml of the flowthrough was retained and
stored at 4°C for cell abundance determination by microscopy (see below).
Determination of cell abundances in the size fractions. The total
number of cells in the unseparated soils, in the pellets resulting from the
Nycodenz gradient, and on each of the filters obtained from each soil was
determined by direct counting procedure on black polycarbonate filters
(0.2-␮m pore diameter) using epifluorescence microscopy and 4,6-diamidino-2-phenylindole (DAPI). For the unfractionated soil samples,
triplicate 1-g subsamples of each soil were suspended in 1 ml of the cell
fixing solution described above, incubated overnight, and vortexed for 15
min. Total bacterial abundances were determined in dilutions from the
unseparated slurries of the soils, from each filtered fraction, and from the
pellets after the cell extraction by incubating aliquots from each diluted
suspension with DAPI at a final concentration of 1 ␮g ml⫺1 for 10 min in
the dark. These solutions were then filtered through a black 0.2-␮m polycarbonate membrane filter. Once completely dried, the filters were
mounted on paraffin oil and cell numbers were determined from at least
20 fields per sample via epifluorescence microscopy. Negative controls
from the cell separation and fractionation processes as well as the solutions used during the dilutions and staining were counted using this approach.
DNA extraction, PCR amplification, and pyrosequencing. Genomic
DNA was extracted directly from soil samples, from the whole-cell fraction separated from the soil particles by Nycodenz gradient, from the
pellet remaining after the Nycodenz gradient procedure, and from the
different-pore-size filters used for the size fractionation. From each of
these 182 individual samples (and the associated negative controls), DNA
was extracted using the MoBio Power Soil DNA extraction kit (MoBio,
Carlsbad, CA) using the modified protocol described by Lauber et al. (17).
A portion of the 16S rRNA gene spanning the V4 hypervariable region was
amplified by triplicate PCRs using the bar-coded primer set (515F/806R),
PCR mixture, and thermal cycling conditions described by Caporaso et al.
(18). This primer set was designed to cover a wide diversity of both Archaea and Bacteria with few biases against individual taxa (19). The replicate PCRs were combined for each sample and quantified using a
PicoGreen double-stranded-DNA (dsDNA) assay (Invitrogen, Carlsbad,
CA). All samples were pooled in equimolar concentrations and sequenced
on a HiSeq2000 instrument (Illumina, San Diego, CA) at The University
of Colorado Advanced Genomics Facility.
Sequence processing. The raw 100-bp DNA sequences generated by
the Illumina HiSeq platform were assigned to their respective samples
using associated bar code sequences and filtered based on quality scores
using the default parameters in QIIME v.1.5.0-dev (20). Because the sequencing generated a large number (19,422,252) of quality-filtered sequences, traditional de novo picking of operational taxonomic units
(OTUs, defined by pairwise sequence similarity) was not practical. Instead, we used an open reference-based OTU picking approach implemented with the QIIME algorithm, “pick_subsampled_reference_otus_
through_otu_table.py,” which allows OTUs to be picked on a practical
timescale while retaining many sequences that are not closely related to
those in published databases. Briefly, this algorithm prefilters sequences
using UCLUST (21) and the Greengenes database preclustered at 97%
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FIG 1 Percentage of total extracted cells from each of the three soils that were recovered within each of the size fractions. Cell counts were determined by direct
microscopy of each individual size fraction, with the size fractions isolated by filtering the Nycodenz-extracted cells through a series of filters of successively
smaller pore sizes. Error bars indicate ⫾1 standard error of the mean.
identity (22) to remove sequences with less than 60% similarity (0.2% of
the sequences). Next, sequences were clustered at the 97% sequence similarity level using the same database. Those sequences which failed to
cluster (26% of the prefiltered sequences) were subsampled for de novo
OTU picking, and representative sequences of these OTUs were subsequently added to a new reference data set. This process was repeated again
with the new reference data set (reference-based OTU picking followed by
de novo picking). A phylogenetic tree was then constructed using sequences aligned by PyNAST (23) in order to calculate phylogenetic dissimilarity matrices, and as a further quality filtering step, OTUs represented by less than 2 sequences and those that could not be aligned were
removed. Taxonomy was assigned to OTUs using the RDP classifier (24)
trained on the Greengenes database set at a 50% confidence threshold.
Following OTU picking, samples were rarefied to a constant sequencing
depth (12,000 sequences per sample) for all downstream analyses. Eighty
samples did not yield a sufficient number of sequences at this rarefaction
depth, so only 102 samples were included in all downstream analyses.
To compare the microbial communities from the unseparated soil
samples, the cell fractions extracted by Nycodenz gradient, and the cells
retained in each fraction size, we used the phylogenetically based unweighted UniFrac metric (25). The resulting distance matrix was used to
determine whether the cell fractions extracted from the different soils
harbored distinct communities by running an analysis of similarity
(ANOSIM) test in PRIMER v6 (26). Likewise, Mantel tests were run in
PRIMER v6 (26) to determine if size fractions more similar in size harbored more similar communities. To describe the size distributions of
individual taxa, we focused on those taxonomic groups (phyla, classes, or
orders) represented by more than 1,000 sequences (summed across all size
fractions per soil sample), as we were not confident in our ability to detect
changes in cell size distributions in taxonomic groups represented by
⬍0.1% of the sequences. The relationships between cell size (determined
by the successive filter pore sizes) and sequence abundances were described by fitting a 3rd-order polynomial function in SigmaPlot (Systat
Software, San Jose, CA).
Nucleotide sequence accession number. All sequence data from this
study have been deposited in the public EMBL-EBI database (http://www
.ebi.ac.uk/) under the accession number ERP003935.
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RESULTS AND DISCUSSION
Cell abundances across size fractions. The three soils (prairie,
forest, and agricultural) harbored numbers of microbial cells that
were within the same order of magnitude (1010 cells g⫺1), with the
forest soil containing the highest cell concentrations, as evident
from the direct cell counts conducted on the unfractionated soil
samples (Table 1). We know from a comparison of cell counts in
the Nycodenz-separated fraction to counts obtained with the unfractionated soils that the majority of the cells were not effectively
separated from the soil matrix using the Nycodenz density gradient approach. We estimate that 13.7%, 2.7%, and 5.2% of the cells
in the prairie, forest, and agricultural untreated soil samples, respectively, were separated from the soil matrix and subsequently
run through the series of size-selecting filters. This efficiency of
cell recovery using the Nycodenz density gradient is low but falls
well within the range of 0.5 to 25% reported in previous studies
that have used a similar approach to separate cells from the soil
matrix (27–30). The variability in cell extraction efficiencies across
the three soils is likely related to differences in soil texture or
organic-matter content (28, 29).
The Nycodenz-separated cells were partitioned into different
size fractions by passage through filters with successively smaller
pore sizes (5, 3, 1.2, 0.8, 0.6, 0.4, and 0.2 ␮m), and direct cell
counts were performed on each fraction (Fig. 1). Across all soils,
⬎60% of the cells were smaller than 1.2 ␮m, confirming results
reported previously (4), with the size distribution in cell abundances varying across different soils. A higher percentage of the
extracted cells from the prairie and forest soil were in the smaller
size classes than in the agricultural soil, where cells were slightly
larger on average (Fig. 1). With only three soils included in this
study, we cannot determine what site or edaphic characteristics
are responsible for the observed differences in size distributions
shown in Fig. 1, but previous work has suggested that such vari-
Applied and Environmental Microbiology
Size Distributions of Soil Bacterial and Archaeal Taxa
FIG 2 Relative abundances of bacterial and archaeal phyla in the bulk soil, i.e., those communities found in the soil prior to Nycodenz extraction (A) and the
relative abundances of phyla in the cell fraction extracted by the Nycodenz gradient (B). Additional details on the taxon abundances are available in Tables S1,
S2, and S3 in the supplemental material for the prairie, forest, and agricultural soils, respectively.
ability in cell size distributions could be related to the taxonomic
structure of the communities, resource availability, or metabolic
state of the microbial cells (10, 14, 27).
Community composition across the studied soils. The microbial communities in the unfractionated samples and the Nycodenz-extracted cells from each of the three soil samples were all
dominated by members of the Proteobacteria, Acidobacteria, Firmicutes, Bacteroidetes, Verrucomicrobia, and Actinobacteria phyla
(Fig. 2; see also Tables S1 to S3 in the supplemental material),
bacterial phyla that are typically the most abundant taxa recovered
from molecular surveys of soil microbial communities (17). However, the microbial communities in each of the three soils (prairie,
December 2013 Volume 79 Number 24
forest, and agricultural) were all significantly different from one
another, both before and after the Nycodenz extraction (Fig. 2)
(ANOSIM, r ⫽ 0.87 and P ⬍ 0.001). Because these soils harbored
distinct communities, the analysis of the microbial taxa found
within each size fraction was conducted separately for each soil.
The Nycodenz separation not only failed to remove all cells
from the soil matrix, as described above, but also failed to extract
all taxa equally. Although the degrees of taxonomic richness
(numbers of OTUs per sample at the rarified sequencing depth)
were similar in the Nycodenz-extracted and unextracted samples
(P ⬎ 0.05 in all cases), community compositions were significantly different between the Nycodenz-extracted and unextracted
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FIG 3 Relative abundances of microbial phyla across the studied size fractions in each soil for those phyla representing more than 2% of the classified sequences
from filter samples. The symbols representing sequence abundances are blue triangles for prairie samples, red circles for forest samples, and green squares for
agricultural samples. The trend line and regression coefficient (r2) are indicated for a specific phylum and soil when the r2 was ⬎0.5. These results were based on
all samples compared at the same sequencing depth (12,000 sequences per sample).
samples for each soil (ANOSIM, r ⬎ 0.6 and P ⬍ 0.03 for all three
soil types). Certain taxa, including members of the Acidobacteria,
Bacteroidetes, and Verrucomicrobia phyla, were significantly overrepresented in the cell fraction extracted by Nycodenz versus the
unextracted soil. In contrast, Firmicutes and Actinobacteria were
underrepresented in the Nycodenz extraction from each soil (Fig.
2). This taxonomic bias introduced during the Nycodenz cell extraction process has been noted previously (31) and may be related, in part, to the differential removal of spore-forming bacteria
during the Nycodenz extraction process (spores have a higher
density than vegetative cells [32]) or due to the tendency of some
taxa to be more closely bound to mineral soil particles than other
taxa (33).
Although the Nycodenz-separated cells represent a biased subset of the cells in the unextracted soils, the same major taxa were
recovered in both the separated and unextracted (whole soil)
communities (Fig. 2). More importantly, despite these taxonomic
biases, we can still determine whether different cell size fractions
harbor distinct bacterial and archaeal communities, and we can
assess the cell size distributions of major bacterial and archaeal
groups. As many approaches used to determine cell size, including
many microscopy-based approaches and the size-fractionation
approach used in this study, require that cells be separated from
the soil matrix, the bias associated with the Nycodenz approach is
somewhat unavoidable until better methods for isolating cells
from the soil matrix are developed.
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Differences in bacterial communities across size fractions.
Microbial community compositions of the different size fractions
retained on the filters were significantly different for the three soil
samples (ANOSIM, prairie, r ⫽ 0.6; forest, r ⫽ 0.23; and agricultural, r ⫽ 0.76; P ⬍ 0.001 at all cases). Moreover, size fractions
closer in size harbored more similar communities than we have
expected by chance, as determined by Mantel tests (prairie, r ⫽
0.66 and P ⬍ 0.001; forest, r ⫽ 0.33 and P ⬍ 0.01; and agricultural,
r ⫽ 0.70 and P ⬍ 0.001). We know of no comparable studies
demonstrating that the communities found in different cell size
fractions are distinct in composition; however, our results are
qualitatively similar to those reported previously (27) where it was
shown the phospholipid fatty acid signatures of cells smaller than
0.4 ␮m were distinct from those found in bulk soil. Our findings
not only highlight that some taxa are often consistently smaller or
larger than other taxa, a point discussed in more detail below, but
also suggest that the soil microbial community is not a homogeneous entity; that subset of a given community composed of
smaller cells will likely interact with the surrounding soil environment in a very different way from the subset with larger cells. For
example, microbial taxa of different sizes are likely to differ in their
nutrient and energy requirements, their abilities to access smallersized soil pores, or their abilities to avoid predation (33–35).
Figures 3 and 4 show the size distributions of major bacterial
and archaeal taxa in each of the soils. Since the three soil types
harbored such distinct communities (Fig. 2), the relative abun-
Applied and Environmental Microbiology
Size Distributions of Soil Bacterial and Archaeal Taxa
FIG 4 Relative abundances of microbial phyla across the studied size fractions in each soil for those phyla representing less than 2% (on average) of the classified
sequences from filter samples. This figure is identical to Fig. 3 except that it shows the less abundant phyla. The symbols representing the sequence abundances
arethe same as for Fig. 3. The trend line and regression coefficient (r2) are indicated for a specific phylum and soil.
dance of individual taxa across the size fractions was dependent on
the soil type in question. Likewise, some taxa were only sufficiently
abundant in one of the three soil types to accurately determine
their size distributions (Fig. 3 and 4). Nevertheless, key patterns
emerge highlighting that those bacteria and archaea common in
soil often show distinct size distribution patterns, with few taxa
being equally abundant in all size classes. For example, at the phylum level we found that Proteobacteria and Actinobacteria were
often relatively more abundant in the larger size fractions (⬎3
␮m), while other phyla, including Acidobacteria, AD3, Verrucomicrobia, and Gemmatimondetes, were consistently more abundant
in the smaller size fractions (⬍0.8 ␮m), with Chloroflexi and Firmicutes being most abundant in the intermediate size fractions.
Most of the rare phyla (those representing ⬍2% of the sequences
for a given sample) and those phyla represented by few (if any)
cultured representatives were most abundant in the smaller size
fractions, including Crenarchaeota, Elusimicrobia, TM6, SC3,
TM7, ZB2, and WS3 (Fig. 4). However, we reiterate that these
phylum-level size distributions were dependent on the soil and
cell size cannot always be predicted from the phylum-level designation.
To determine if the differences in the size distributions of individual phyla observed across the soils were related to the predominance of different subphyla in the three soils, we investigated
size distributions at finer levels of taxonomic resolution for some
of the more abundant phyla (see Fig. S1 in the supplemental material). We found that different groups within a given phylum
December 2013 Volume 79 Number 24
often exhibited distinct size distribution patterns. For example,
some actinobacterial orders (including Actinomycetales) had consistently larger cell sizes than other orders within this phylum (see
Fig. S1), patterns that were not obvious at the phylum level of
resolution (Fig. 3), with similar patterns observed for individual
orders within the Proteobacteria and Bacteroidetes phyla (see Fig.
S1). Clearly, different taxonomic groups often have distinct cell
sizes, and these patterns are evident at various levels of taxonomic
resolution.
What does cell size tell us about microbial life history strategies? There are two possible reasons why many of the soil microbial taxa shown in Fig. 3 and 4 exhibit uneven distributions across
the cell size fractions. First, some taxa may be intrinsically larger
than other taxa, regardless of growth conditions, resource availability, or growth phase. For example, many actinobacterial taxa
are known to have a filamentous growth form that would effectively give them a larger cell size than nonfilamentous bacteria
regardless of the specific growth conditions. Alternatively, cell
morphology may be a more plastic trait, determined not by the cell
genotype but with cell size varying as a function of growth conditions or growth phase. We know that this is true for many cultured
taxa that are often smaller when in the stationary phase of growth
(36) or when carbon and nutrient resources are limiting (10). In
this study, we could not differentiate between these two possible
explanations, and in all likelihood, both factors may contribute to
the variation in cell size observed across the soil taxa examined. A
similar point has been made previously where it has been hypoth-
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TABLE 2 Size classifications of the phyla that were reasonably abundant
in the soils examined (represented by more than 1,000 sequences), the
number of sequences deposited in the Ribosomal Database Project
(RDP [43]) for each of these phyla, and the number of these sequences
that were obtained from bacterial or archaeal isolatesa
Phylum
Size
No. of
Total no. of sequences
sequences from isolates
in the RDP in the RDP
Actinobacteria
Cyanobacteria
Proteobacteria
Firmicutes
Planctomycetes
Chloroflexi
Acidobacteria
Gemmatimonadetes
Crenarchaeota
Verrucomicrobia
Elusimicrobia
TM7
Large
Large
Large
Intermediate
Intermediate
Intermediate
Small
Small
Small
Small
Small
Small
180,827
21,330
341,501
420,503
11,233
20,447
14,151
1,256
7,123
9,586
172
2,167
32,101
3,851
89,411
54,291
416
179
165
7
252
158
3
12
% of RDP
sequences
represented
by isolates
17.7
18.1
26.2
12.9
3.7
0.8
1.2
0.6
3.5
1.6
1.7
0.6
a
Cells were considered large, intermediate, or small if they were in the range of 5 to 3
␮m, 3 to 0.8 ␮m, or 0.8 to 0.2 ␮m, respectively (see Fig. 3 and 4). Only those taxa that
exhibited a discernible peak in relative abundances within a given size range in any of
the soils were included here. Candidate divisions not recognized in the RDP database
are not listed.
esized that the smaller microorganisms either may represent distinct groups of intrinsically small taxa or may be starved, dormant
forms of bigger microbes (10, 37, 38).
Despite the uncertainties in delineating the specific factors
driving variation in cell size across taxa, some interesting patterns
emerge when we take the data presented in Fig. 3 and 4 and divide
the taxa into general size classes (Table 2), as most of the smaller
taxa were those with few cultivated representatives. This general
pattern suggests that cell size is an important ecological trait, with
smaller taxa having attributes that make them more difficult to
culture. A similar pattern was also noted previously (6, 11) where
it was shown that novel taxa were relatively common in the
smaller cell sizes (⬍0.45 ␮m) and less easily cultivated. The general ecological attributes associated with smaller cells may include
low growth rates, as the presence of fast-growing bacteria would
occlude colony formation of slower-growing taxa, especially on
nutrient-rich media (13, 39). Smaller taxa may also be better
adapted for growth in oligotrophic conditions than larger cells,
and we would expect smaller cells to be underrepresented in culture-based surveys of microbial diversity unless culture conditions and media are designed to select for oligotrophic taxa (e.g.,
see references 40 to 42). Interestingly, those soil bacteria that were
most readily cultivated under oligotrophic conditions, including
members of the Acidobacteria, Gemmatimonadetes, and Verrucomicrobia phyla (40, 41), were also those phyla that tended to be in
the smaller cell size fractions (Fig. 3).
In soil, the ecological attributes associated with a smaller cell
size may be advantageous under certain conditions, including
more efficient nutrient uptake when resources are limiting due to
a larger surface area-to-volume ratio, increased protection against
predation, and the ability to occupy microenvironments different
from those occupied by larger cells (3). Efforts to characterize soil
microbial groups that are underrepresented in culture conditions,
using either single-cell isolation or enrichment culturing strate-
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gies, could benefit from first size selecting soil microbial cells, as
many poorly studied groups appear to have cell sizes that are
smaller than average.
Conclusions. Different cell size fractions harbored distinct bacterial and archaeal communities, with many of the taxa exhibiting
unique size distribution patterns (few taxa were equally abundant
across all size classes). The smallest size fractions contained a higher
proportion of rare phyla and candidate divisions (including Acidobacteria, Gemmatimonadetes, Crenarchaeota, Verrucomicrobia, Elusomicrobia, and the candidate divisions AD3, TM6, SC3, TM7, ZB2,
and WS3) than the larger size fractions. However, soil type was an
important factor influencing both the taxa found within different size
fractions and the total abundance of cells in each size fraction, highlighting that the size distribution pattern for a given taxonomic group
is not constant and can vary depending on soil properties and the
members of the taxonomic group. As suspected, cell size is an important ecological trait, and by combining cell size fractionation with
other approaches (including genomics-based approaches), we can
better understand the ecology and life history strategies of the vast
majority of soil microbial taxa that remain poorly characterized.
ACKNOWLEDGMENTS
We thank Jessica Henley and other members of the Fierer laboratory for
their assistance with this project.
M.C.P. received funding for this work from both the Fulbright Program and the Spanish Ministry of Education through the National Program of Mobility and Human Resources from the National Plan I-D⫹I
2008-2011. N.F., C.L.L., and J.W.L. were supported by grants from the
National Science Foundation (DEB-0953331) and the U.S. Department of
Agriculture.
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