APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Sept. 2005, p. 5404–5410
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
Vol. 71, No. 9
Extracellular DNA in Single- and Multiple-Species
R. E. Steinberger and P. A. Holden*
Donald Bren School of Environmental Science and Management, University of California,
Santa Barbara, California 93106
Received 1 September 2004/Accepted 29 March 2005
The extracellular polymeric substances (EPS) of bacterial biofilms form a hydrated barrier between cells and
their external environment. Better characterization of EPS could be useful in understanding biofilm physiol-
ogy. The EPS are chemically complex, changing with both bacterial strain and culture conditions. Previously,
we reported that Pseudomonas aeruginosa unsaturated biofilm EPS contains large amounts of extracellular
DNA (eDNA) (R. E. Steinberger, A. R. Allen, H. G. Hansma, and P. A. Holden, Microb. Ecol. 43:416–423, 2002).
Here, we investigated the compositional similarity of eDNA to cellular DNA, the relative quantity of eDNA, and
the terminal restriction fragment length polymorphism (TRFLP) community profile of eDNA in multiple-
species biofilms. By randomly amplified polymorphic DNA analysis, cellular DNA and eDNA appear identical
for P. aeruginosa biofilms. Significantly more eDNA was produced in P. aeruginosa and Pseudomonas putida
biofilms than in Rhodococcus erythropolis or Variovorax paradoxus biofilms. While the amount of eDNA in
dual-species biofilms was of the same order of magnitude as that of of single-species biofilms, the amounts were
not predictable from single-strain measurements. By the Shannon diversity index and principle components
analysis of TRFLP profiles generated from 16S rRNA genes, eDNA of four-species biofilms differed signifi-
cantly from either cellular or total DNA of the same biofilm. However, total DNA- and cellular DNA-based
TRFLP analyses of this biofilm community yielded identical results. We conclude that extracellular DNA
production in unsaturated biofilms is species dependent and that the phylogenetic information contained in
this DNA pool is quantifiable and distinct from either total or cellular DNA.
mixture of hydrated polymers that serve various purposes (59),
including nutrient (62) and water (46) retention and protection
from toxins such as antibiotics (52, 53) and pollutants (61). EPS is
often described as polysaccharide, although the model organism
in biofilm studies, Pseudomonas aeruginosa, has long been known
to excrete large amounts of DNA (19). Extracellular DNA
(eDNA) has previously been shown to be essential for saturated
biofilm stability during the early stages of biofilm growth (58). We
recently reported that extracellular DNA was continuously
present in unsaturated Pseudomonas aeruginosa biofilms (51) and
was maximally 50% more abundant than cellular DNA. Since
unsaturated biofilms are not subjected to hydrodynamic shear,
the structural roles for eDNA in this context are questionable.
The eDNA may also enhance gene transfer (36) and provide
known about the universality, composition, and persistence of
eDNA, particularly in natural and multiple-species environments,
so little more than speculation about its purpose is possible at the
moment. These concerns should be addressed to improve our
understanding of biofilm physiology; they also could have impli-
Culture-independent techniques have allowed scientists to
catalog and compare microbial communities of such diverse
natural environments as oligotrophic waters (11, 24), marine
and freshwater sediments (57, 60), and surface and subsurface
soils (2, 10, 13, 25, 27, 30), as well as artificial environments
such as water distribution pipes and sewage treatment plants
(12, 20, 48). The most commonly used culture-independent
techniques for microbial community analysis depend on the
isolation and amplification via PCR of conserved genes, such
as those encoding 16S rRNA (15). Two approaches to isolating
microbial community DNA from environmental samples were
developed almost 20 years ago—the direct extraction approach
pioneered by Ogram et al. (38) and the indirect extraction
approach pioneered by Holben et al. (21)—but debate as to
which is superior continues today (17, 45). The direct extrac-
tion approach is often preferred because it generally yields
more DNA with fewer steps (17). However, this approach will
also extract eDNA along with cellular DNA. The original pro-
cedure of Ogram et al. (38) included extra washing steps to
remove the eDNA prior to cell lysis, and some investigators
continue to consider eDNA (for examples, see references 2, 16,
33, and 55) in their extraction protocol. However, others (for
examples, see references 27, 30, 47, and 63) don’t, which could
be attributed to the widespread assumption that eDNA in
natural, complex environments is of low abundance (37).
If little eDNA is produced and degradation is rapid, then the
amount of eDNA found in natural environments will be small.
However, soil concentrations of eDNA are as high as 2 ?g/g of
dry soil (37), and eDNA can comprise more than 70% of the
total DNA pool in marine sediments (9). The actual amount of
eDNA may be even higher since the majority of eDNAs may
not be released even with multiple sequential extractions (29).
While eukaryotic DNA (18) and dissolved DNA (42) are
quickly degraded by environmental nucleases, bacterial DNA
(2) and recombinant genes of bacterial origin (40) may persist
* Corresponding author. Mailing address: Bren Hall, UCSB, Santa
Barbara, CA 93106-5131. Phone: (805) 893-3195. Fax: (805) 893-7612.
outside cells as PCR-amplifiable fragments for months or
longer. Binding to sediments (1, 9, 29), clays (3), and humic
acids (8) can protect eDNA from degradation by nucleases,
preserving its ability to transform cells (1, 8) and be amplified
by PCR (2, 3). The DNA binding appears to result from in-
teraction with a large number of low-specificity binding sites
(43) which favors the retention of large DNA fragments. Ex-
traction of only eDNA yields a unique community profile quite
different from that of bacterial cells extracted from the same
In this paper, we assayed single- and dual-species bacterial
biofilms for relative eDNA content to determine the univer-
sality of eDNA production and the persistence of eDNA in
dual-species biofilms. To gain insight into the composition of
eDNA, we compared eDNA with cellular DNA isolated from
P. aeruginosa using randomly amplified polymorphic DNA
(RAPD). Finally, we investigated the proportional effects of
eDNA versus cellular DNA on culture-independent assess-
ments of microbial community composition and diversity. Our
findings reveal the strain specificity of eDNA abundance and
the unpredictable accumulation of eDNA in dual-species bio-
films. Further, eDNA has a distinct signature of 16S rRNA
genes relative to cellular and total biofilm DNA.
MATERIALS AND METHODS
Bacterial strains and chemicals. Four strains of bacteria were used in cultur-
ing biofilms. P. aeruginosa strain PG201 (Urs Ochsner, University of Colorado)
and Pseudomonas putida strain mt-2 (Gary Sayler, University of Tennessee—
Knoxville) are both gram-negative soil isolates. Rhodococcus erythropolis was
provided by Joe Lepo (University of West Florida) and is a mucoid gram-positive
environmental isolate. Variovorax paradoxus is a gram-negative environmental
isolate (22) that appears to be a vadose zone specialist (22, 25). All strains were
archived in 70% Luria broth–30% glycerol at ?80°C. The specific growth rate
constants of the strains in liquid culture (10% tryptic soy broth [TSB], 30°C, 200
rpm) were 0.387/h for P. aeruginosa, 0.397/h for P. putida, 0.150/h for R. eryth-
ropolis, and 0.197/h for V. paradoxus.
All chemicals, unless otherwise specified, were reagent grade or higher (Sig-
ma-Aldrich, St. Louis, MO).
Biofilm growth and harvesting. Biofilms were cultivated for three purposes: to
quantify the abundance of eDNA, to compare the composition of cellular DNA
to eDNA via RAPD analysis, and to quantify the impact of eDNA on the
[TRFLP]) analysis of a biofilm bacterial community. For eDNA quantification
and TRFLP analysis, biofilms were cultured using the method of Auerbach et al.
(4) on Nuclepore polyester membranes (13-mm diameter, 0.1-?m pore, 6-?m
thickness; Whatman, Clifton, NJ) overlaying 10% tryptic soy agar (3 g; TSB, 15 g
Bacto Agar per liter [both from Difco, Fisher Scientific, Pittsburgh, PA]). To
recover larger quantities of DNA for RAPD analysis, P. aeruginosa strain PG201
was cultured similarly on larger polyester Nuclepore membranes (47-mm diam-
eter, 0.1-?m pore size) over Luria broth agar plates for 5 days. Prior to inocu-
lation, membranes were sterilized (70% ethanol for 2 min), air dried, and then
transferred with sterile forceps to the solid medium surface. Small membranes
were spaced equally in a pentagon formation centered in the petri dish to
facilitate similar zones of nutrient availability during growth. Inocula were pre-
pared from suspensions of single-strain bacterial cultures (10% TSA at 30°C for
3 days) in sterile 0.9% NaCl solution. To facilitate the equal representation of
species in mixed inocula, the densities (by absorbance, 600 nm) of the suspen-
sions were equalized by adjusting them with sterile 0.9% NaCl. For single-species
biofilms, each membrane was inoculated with 4 ?l of suspension onto the center
of each membrane. For multiple-species biofilms, 2 ?l of each inoculum was
deposited onto the center of each membrane into the same droplet so that
dual-species membranes received a total of 4 ?l of inoculum and biofilms com-
posed of all four species received a total of 8 ?l of inoculum. As with previous
experiments, the inocula comprised less than 1% of the final assayed biofilm (50).
Uninoculated sterile controls were prepared in the same manner. The petri
dishes were sealed with Parafilm to prevent drying and incubated at 30°C in the
dark. Biofilms grow radially and concentrically away from the point of inocula-
After 120 h, three membranes were harvested for each species combination
and for the sterile control. The process of biofilm harvesting and EPS separation
was based on the method proposed by May and Chakrabarty (35). Membranes
were lifted from the agar surface with biofilm intact using sterile forceps. Each
biofilm was dispersed by vortexing the membrane in 1 ml of 0.9% NaCl solution
containing 10 mM EDTA for small membranes and 3 ml of 0.9% NaCl–10 mM
EDTA solution for large membranes. The 0.9% NaCl has a lower water potential
(?0.71 MPa) than the growth medium (?0 MPa) to prevent cell lysis during
harvesting. EPS and cells were separated by centrifugation (12,000 ? g for 30 min
at 4°C) and stored at ?20°C until analysis. Based on the size of the P. aeruginosa
genome (6.3 Mbp ) and the number of cells remaining in the biofilm (no
more than 0.0001%), contaminating cellular DNA comprises approximately
0.01% of the total DNA in the EPS.
DNA extraction, quantification, and purification. The thawed cell pellets of
the single- and dual-species biofilms were resuspended in 500 ?l of 1 N NaOH
and heated at 80°C for 1 h to complete lysis. The NaOH was then neutralized
with 500 ?l of 1 N HCl. For RAPD and TRFLP, DNA was released from thawed
cell pellets by bead beating in sterile 0.9% NaCl for 30 s using 0.5 g of 100-?m
sterile glass beads to prevent possible sequence degradation. The DNA in the
lysates and the eDNA in the supernatant were quantified using Picogreen flu-
orometry (Molecular Probes, Eugene, OR) with calf thymus DNA as a standard.
DNA was quantified prior to purification because of possible purification bias.
Both cellular DNA and eDNA were purified and concentrated by extracting
twice with an equal volume of 25:24:1 phenol-chloroform-isoamyl alcohol and
then precipitating with 1/10 volume of 3 M potassium acetate and 2/3 volume of
isopropanol. After centrifugation (10 min, 4°C, 10,000 ? g), the pellet was
washed with 100% ethanol and air dried. The dried DNA pellet was resuspended
in 20 ?l of 10 mM Tris-Cl, pH 8.5. The concentration and purity of the purified
DNA were determined spectrophotometrically by the absorbance ratio A260/
A280. Purified DNA solutions were adjusted to 25 ng/ml with 10 mM Tris-Cl, pH
RAPD analysis of P. aeruginosa DNA. To infer the composition of eDNA,
RAPD was performed on P. aeruginosa cellular and eDNA according to the
method of Mahenthiralingam et al. (32). Eight different 10-mer primers (Table
1) previously shown to produce reproducible and distinguishable RAPD profiles
with P. aeruginosa (32) were obtained from Qiagen (Valencia, CA). Each 25-?l
reaction contained 40 ng of genomic DNA, 40 pmol of oligonucleotide, 1 U Taq
polymerase (Qiagen), 250 ?M (each) deoxynucleoside triphosphates (Qiagen), 3
mM MgCl2, and 5 ?l Q solution (Qiagen) in PCR buffer (Qiagen). Amplification
was done in a PCRSprint thermocycler (Hybaid US, Franklin, MA) as follows: (i)
4 cycles, each consisting of 5 min at 94°C, 5 min at 36°C, and 5 min at 72°C, (ii)
30 cycles, each consisting of 1 min at 94°C, 1 min at 36°C, and 2 min at 72°C, and
(iii) a final extension step of 10 min at 72°C.
The RAPD products were separated by gel electrophoresis on a 2% agarose
gel in 0.5% Tris-borate-EDTA using a Mini-Sub Cell GT system (Bio-Rad,
Hercules, CA) at 100 V (approximately 25 mA) for 45 min. The gel was subse-
quently stained with ethidium bromide and photodocumented. Band intensity
was quantified using ImageQuant software (Molecular Dynamics, Sunnyvale,
CA). Pairwise comparisons of lanes were performed based on the presence or
absence of bands, as identified by the ImageQuant software. Comparisons were
also made by comparing the brightness along linear paths of equal length in each
lane. The intensities were plotted against the location, and the correlation coef-
ficient was calculated using the correlation tool of the Data Analysis ToolPak in
Microsoft Excel 2000.
TRFLP analysis. 16S rRNA genes were amplified from purified DNA using
eubacterial primers 8F Hex (fluorescently labeled 5?AGAGTTTGATCCTGGC
TABLE 1. Sequences of 10-mer RAPD primersa
Primer name Sequence (5? to 3?)
aData from reference 31.
VOL. 71, 2005EXTRACELLULAR DNA IN UNSATURATED BIOFILMS5405
TCAG ) and 1389R (5?ACGGGCGGTGTACAAG ). Each 100-?l reaction
contained 75 ng template, 2.5 U Taq polymerase, 200 ?M (each) deoxynucleoside
triphosphates, 50 pmol DNA primers, 2.2 mM MgCl2(including 1.5 mM from the
reaction buffer), 1? reaction buffer and Q mix (Qiagen, Valencia, CA), and 2 mg/ml
bovine serum albumin (Promega, Madison, WI). Amplification was done in a
PCRSprint thermocycler (Hybaid US) according to the following hot-start protocol:
(i) denaturing for 4 min at 95°C, extension for 1 min at 58°C, and annealing for 90 s
at 58°C, and 90 s at 72°C, and (iii) a final elongation for 10 min and 45 s at 72°C.
After amplification, PCR product was purified using QIAquick PCR purification
columns (Qiagen, Valencia, CA). Amplicons were then restricted using HhaI (New
England Biolabs, Beverly, MA) in a 20-?l reaction containing 300 ng purified PCR
product, 1 U HhaI enzyme, 1? reaction buffer, and 0.1 mg/ml bovine serum albumin
overnight at 37°C. Following restriction, the enzyme was denatured (65°C for 20
min). Samples were desalted using QIAquick nucleotide removal columns (Qiagen,
Valencia, CA) and eluted into distilled water. The lengths of the fluorescently
labeled fragments were determined with an Applied Biosystems Instruments (Foster
City, CA) Prism 3100 genetic analyzer. Peak areas for replicated peaks were nor-
malized by total replicated peak area, and unreplicated peaks were not included in
subsequent data analysis. Shannon-Weaver diversity and evenness indices were cal-
culated according to the method of Brodie et al. (7). Principal components analysis
(PCA) and correspondence analysis of shared peaks were performed using PC-Ord
(MjM Software Design, Gleneden Beach, OR). Both types of ordination were
performed to verify that problems such as normalization bias (23) and the horseshoe
effect (41) were minimized. In order to assess variability in the TRFLP profiles,
independent replicates for total DNA (n ? 8), cellular DNA (n ? 7), and eDNA (n
? 4) were conducted from the PCR step forward. Due to the similarity of the total
DNA and cellular DNA profiles, TRFLP analysis was replicated twice more for
these two pools to strengthen the statistical significance of their interpretation.
Production and persistence of eDNA. Four species (P.
aeruginosa, P. putida, R. erythropolis, and V. paradoxus) were
tested for the ability to produce eDNA in single- and dual-
species biofilms. Both strains of Pseudomonas produced high
levels of eDNA, averaging 0.22 ? 0.04 g of eDNA/g of cellular
DNA for P. aeruginosa and 0.51 ? 0.10 g of eDNA/g of cellular
DNA for P. putida (Fig. 1A), although P. putida produced
significantly more (P ? 0.017) than P. aeruginosa. Both V.
paradoxus and R. erythropolis produced very small but statisti-
cally similar (P ? 0.10) amounts of eDNA (0.05 ? 0.02 and
0.029 ? 0.006 g of eDNA/g of cellular DNA, respectively)
which was significantly less eDNA (P ? 0.005) than either of
the Pseudomonas species. Of the total DNA in the Pseudomo-
nas biofilms, 17 ? 3% and 32 ? 4% were extracellular for P.
aeruginosa and P. putida, respectively. eDNA represented less
than 5% of the total DNA for R. erythropolis and V. paradoxus.
The levels of eDNA in the dual-species biofilms (relative to
the concentration of cellular DNA) were similar to the levels of
eDNA observed in single-species biofilms (Fig. 1) but were not
necessarily additive or quantitatively predictable from the com-
posite eDNA from individual strains. Combining the two high
eDNA-producing strains (P. aeruginosa and P. putida) led to a
high eDNA content not significantly different (P ? 0.1) from
either P. aeruginosa or P. putida single-species biofilms. Simi-
larly, combining the low eDNA-producing strains (R. eryth-
ropolis and V. paradoxus) produced a biofilm with an eDNA
content statistically similar (P ? 0.01) to the biofilms of the
component strains. However, mixing the high and low eDNA-
producing strains led to unpredictable results. Combining ei-
ther Pseudomonas species with R. erythropolis produced a high
eDNA biofilm (0.36 ? 0.13 g/g of cellular DNA and 0.39 ?
0.09 g/g of cellular DNA for P. aeruginosa and P. putida, re-
FIG. 1. Average eDNA found in single-species biofilms (A) and multiple-species biofilms (B). The eDNA has been normalized to cellular
DNA. The species’ names have been abbreviated as follows: PA, P. aeruginosa; PP, P. putida; RE, R. erythropolis; VP, V. paradoxus. Error bars
represent 1 standard error of each mean. For single-species biofilms, n ? 7. For multiple-species biofilms, n ? 3. Bars with different letters are
significantly different (P ? 0.05).
5406STEINBERGER AND HOLDENAPPL. ENVIRON. MICROBIOL.
spectively) whose eDNA content was not statistically different
from that of either Pseudomonas species (P ? 0.1). However,
combining either Pseudomonas species with V. paradoxus pro-
duced a biofilm with relatively little eDNA (0.07 ? 0.02 g/g of
cellular DNA and 0.14 ? 0.02 g/g of cellular DNA for P.
aeruginosa and P. putida, respectively). The amount of eDNA
in composite biofilms of P. aeruginosa and V. paradoxus was not
significantly different from the single-species biofilms of either
R. erythropolis or V. paradoxus (P ? 0.1), while the eDNA of
composition P. putida and V. paradoxus biofilms was signifi-
cantly different from both (P ? 0.01). It appears that while the
bacterial species composing the biofilm do affect the amount of
eDNA in a biofilm, they don’t do so in a predictable manner.
RAPD comparison of P. aeruginosa cellular DNA and eDNA.
The eDNA and cellular DNA pools of P. aeruginosa unsatur-
ated biofilm were compared using the technique of RAPD
analysis and eight different primers (Table 1). For all primers,
the bright bands were common to the cellular DNA and eDNA
(Fig. 2). Some of the fainter bands, particularly those with
higher molecular weight, were present only in either cellular
DNA or eDNA. These variations, however, were similar to the
level of variations observed between replicate RAPD reactions
(Fig. 2B). The correlation coefficients, based on band bright-
ness values from cellular DNA and eDNA RAPDs, ranged
from 0.790 to 0.989 (average, 0.913) and were equivalent to
those of replicate RAPDs (0.823 to 0.992; average, 0.934). The
eDNA appeared to produce brighter, more consistent bands of
higher-molecular-weight fragments, while the cellular DNA
produced more consistent bands of lower-molecular-weight
fragments (Fig. 2).
TRFLP analyses of cellular DNA, eDNA, and total DNA. We
compared the TRFLP profiles generated from cellular DNA,
eDNA, and total DNA extracted from a biofilm containing all
four species of bacteria. The resulting TRFLPs (Fig. 3) showed
three major peaks and six minor peaks. The major peaks,
located at 61 bp, 152 bp, and 566 bp, appeared in all the
TRFLPs, as did the minor peaks located at 56 bp, 202 bp, 205
bp, and 438 bp. The remaining minor peaks, located at 140 bp
and 562 bp, appeared only in the cellular DNA and the total
DNA. Replication of TRFLP profiles for each DNA source (n
? 4) showed excellent reproducibility, with the peak areas of
each fragment length having variances ranging from 2.5% to
29.1% (average, 11.7%).
The TRFLP profile of the total DNA appeared very similar
to that of the cellular DNA but slightly different from that of
the eDNA (Fig. 3). Individual peak areas of the eDNA TRFLP
fragments were significantly different (P ? 0.05) from those of
the cellular DNA profiles for all but the 56-bp fragment (Fig.
4). The peak areas of fragments in the total DNA TRFLP
profiles were significantly different (P ? 0.05) from those of the
eDNA TRFLP profiles but were not significantly different (P ?
0.1) from those of the cellular DNA TRFLP profiles (Fig. 4).
The average Shannon-Weaver diversity indices for the total
DNA, cellular DNA, and eDNA TRFLP profiles were 1.606 ?
0.041, 1.647 ? 0.021, and 1.244 ? 0.050, respectively. The
average Shannon-Weaver index of the eDNA profile was sig-
nificantly lower (P ? 0.001) than either the cellular DNA or
total DNA profiles. The average Shannon-Weaver diversity
index from the profile of total DNA was statistically similar (P
? 0.1) to that of the cellular DNA profile. The average even-
ness indices for the total DNA, cellular DNA, and eDNA
TRFLP profiles were 0.732 ? 0.018, 0.750 ? 0.009, and 0.640
? 0.026, respectively, and followed the same statistical pattern.
The PCA of the TRFLP profiles was able to explain 66% and
24% of the variability between replicates on the first and sec-
ond axes, respectively (Fig. 5). The replicates from the cellular
FIG. 2. Comparative RAPD analysis of P. aeruginosa cellular DNA
and eDNA. (A) Three replicate cellular DNA and three replicate
eDNA amplification products for primers 270 (left of the ladder) and
275 (right of the ladder) were compared to ascertain banding pattern
reproducibility. Center lane, 1-kb ladder. (B) RAPDs of cellular DNA
(left lane in each pair) and eDNA (right lane in each pair) for primers
270, 272, 275, 277, and 287, from left to right. The lanes are labeled as
follows: C, cellular DNA; E, eDNA; and L, ladder.
FIG. 3. TRFLP profiles generated from cellular DNA (top), eDNA
(middle), and total DNA (bottom) from a multiple-species unsatur-
ated biofilm comprised of P. aeruginosa, P. putida, R. erythropolis, and
VOL. 71, 2005EXTRACELLULAR DNA IN UNSATURATED BIOFILMS5407
DNA and eDNA TRFLPs were grouped closely and clearly
separated from each other. The total DNA TRFLP profiles
also grouped, though less tightly, and overlapped the cellular
DNA profiles. The ordination by correspondence analysis pro-
duced very similar distribution and groupings to those of the
PCA ordination (data not shown).
Previously we reported that the EPS of unsaturated bacterial
biofilms has a complex chemistry, comprised of eDNA, poly-
saccharides, and protein (51). Here, we investigated species
composition as a factor governing eDNA and thus EPS com-
plexity; we also investigated the potential for eDNA to influ-
ence culture-independent microbial community analyses. Our
experimental results lead to several conclusions. First, some
but not all bacteria produce large amounts of eDNA that
persist when grown as either single- or dual-species unsatur-
ated biofilms. Second, even if an isolated species is not a large
producer of eDNA, the presence of a low eDNA producer can
significantly influence the amount of eDNA in multiple-species
biofilms. One explanation of this conclusion might be compet-
itive interactions between strains that ultimately favor the
growth of the lower eDNA producer (e.g., V. paradoxus). How-
ever, other explanations are also possible including ecological
interactions due to differing growth rates (see Methods), dif-
ferent rates of cell death and lysis (26, 56), and the strain-
dependent utilization of eDNA as a nutrient (14). Our obser-
vation that the genetic composition of eDNA and cellular
DNA differed significantly in multispecies biofilms suggests
that the mechanisms are likely to be complex and deserve
Another conclusion concerns the possible role of eDNA in
confounding the analysis of natural microbial communities.
Here, we report that the primary sequence of eDNA cannot be
distinguished from that of cellular DNA for some species.
Although eDNA did not contribute to the overall TRFLP
profile in our low-diversity biofilms, eDNA and cellular DNA
profiles differed significantly by TRFLP. Given that eDNA is
typically recovered during DNA extraction (for examples, see
references 2 and 38) and that it accumulates in biofilms dis-
proportionately to the initial species distribution, eDNA could
affect the culture-independent assessments of communities.
Community diversity, ambient conditions, and mineralogy
(e.g., clay content) are just a few of the variables in natural
environments that could affect how eDNA affects the culture-
independent assessments of whole communities. Previously,
16S rRNA gene amplicons in eDNA were shown to form a
distinct denaturing gradient gel electrophoresis (DGGE) pro-
file (2), but eDNA contributions to the total DGGE profile
were not reported. Furthermore, our comparison of profiles
was limited to that generated by amplifying genes for small-
subunit rRNA. Given the amount of DNA outside of the cells,
we cannot rule out that other sequences of interest, i.e., some
encoding catabolic functions, could be amplified dispropor-
tionately from eDNA in comparison to biofilm species com-
position. Thus, because both the relative proportions of eDNA
and cellular DNA in natural environments and the effect of
eDNA on other nucleic acid-based assays (31) are currently
unknown, it seems premature to dismiss any possible effect of
eDNA on current culture-independent assessments.
Several questions still remain about the structure of eDNA.
For example, why would more higher-molecular-weight RAPD
products appear for P. aeruginosa eDNA relative to cellular
DNA? One explanation could be disproportionate shearing of
cellular DNA during the lysis of P. aeruginosa cells. However,
the higher intensity could also be due to a difference in the
average molecular weight of eDNA or to a difference in mo-
lecular conformation (i.e., secondary/tertiary structure.) While
eDNA appears to have the same primary sequence as cellular
DNA, whether eDNA is circular or linear and what proteins (if
any) are associated with the secondary structure and mainte-
nance of the eDNA are still unknown. eDNA of environmental
biofilms may also be selectively enriched in specific sequences,
particularly since coculturing microbes has been shown to stim-
ulate plasmid DNA release (34). The resistance of biofilms to
destruction by DNase after an initial growth period (58) may
suggest that the DNA in the biofilm is somehow stabilized,
which may inhibit the effectiveness of proposed biofilm treat-
ment methods involving DNase (44). A better understanding
of the structure of the eDNA may also elucidate why it con-
FIG. 4. Average percentage of total peak area contributed by indi-
vidual fragments to the TRFLP profiles of cellular DNA (n ? 7), total
DNA (n ? 8), and eDNA (n ? 4). Error bars represent the standard
error of each mean.
FIG. 5. Plot of principle components 1 (PC1) and 2 (PC2) repre-
senting the percent variability (axis labels) of the TRFLP profiles from
cellular DNA, eDNA, and total DNA. PCA was performed from a
similarity matrix based on comparing peak areas.
5408 STEINBERGER AND HOLDENAPPL. ENVIRON. MICROBIOL.
tributes so little to the total community fingerprint when ana-
lyzed by DGGE (2) or TRFLP.
In addition to the structure of eDNA, there are also many
questions about the origins and functions of eDNA. It is un-
clear whether eDNA is excreted, as suggested by Hara and
Ueda (19), or is a product of cell lysis, a natural part of biofilm
development (26, 56). The variable levels of eDNA produced
among bacterial species suggests that eDNA may be an envi-
ronmental adaptation, although the ecological importance re-
mains unknown. Strong cases have been made for the roles of
eDNA in exchanging genetic information (14, 34, 36), in sta-
bilizing biofilms (36, 58), and as a nutrient during starvation
(14). It is probable that eDNA also fulfills many roles fre-
quently ascribed to exopolysaccharides (59). Continued study
of eDNA is warranted to eliminate uncertainty about the ef-
fects on apparent microbial community composition by cul-
ture-independent techniques and to discern its roles in the
microbial ecology of diverse environments. More broadly, a
better understanding of eDNA may have consequences beyond
its effect on analyses of community composition. For example,
more knowledge of eDNA structure could allow researchers to
increase the effectiveness of DNase treatments for cystic fibro-
sis and persistent infections (5, 6). Further, because the
amount of eDNA in EPS can substantially impact the physio-
chemical properties of industrial biofilms (49), knowing the
origin and function of eDNA in saturated biofilms could be of
economic importance. Finally, the cryptic sequences in eDNA
could potentially be used to trace the growth and transport of
bacteria in natural environments (2).
We thank Gretchen Hoffman from the Department of Ecology,
Evolution, and Marine Biology, University of California—Santa Bar-
bara, for the use of ImageQuant software and equipment.
This work was supported by U.S. EPA STAR Fellowship
U-91583201 to R.E.S. and by the NSF Ecology and Microbial Obser-
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