JOURNAL OF BACTERIOLOGY, Mar. 2007, p. 2125–2127
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
Vol. 189, No. 5
Possible Nonconductive Role of Geobacter sulfurreducens Pilus
Nanowires in Biofilm Formation?
Gemma Reguera,* Rachael B. Pollina,† Julie S. Nicoll,‡ and Derek R. Lovley
Department of Microbiology, University of Massachusetts, Amherst, Massachusetts 01003
Received 14 August 2006/Accepted 29 November 2006
Geobacter sulfurreducens required expression of electrically conductive pili to form biofilms on Fe(III) oxide
surfaces, but pili were also essential for biofilm development on plain glass when fumarate was the sole electron
acceptor. Furthermore, pili were needed for cell aggregation in agglutination studies. These results suggest that
the pili of G. sulfurreducens also have a structural role in biofilm formation.
One of the hallmarks of Geobacter species is their ability to
conserve energy from the transfer of electrons to a variety of
extracellular electron acceptors, such as metals [Fe(III),
Mn(IV), and U(VI)], humic acids, and electrodes (5, 6). Es-
tablishing an electrical connection with an extracellular elec-
tron acceptor poses challenges not faced by microorganisms
that reduce soluble electron acceptors within the cell. In con-
trast to other Fe(III) oxide-reducing bacteria (4, 9–11), such as
Shewanella and Geothrix species, Geobacter species do not ex-
crete electron shuttles (8) and require direct contact with the
electron-accepting surface (1, 10). Previous studies (14) have
demonstrated that the pili of Geobacter sulfurreducens are con-
ductive and that expression of pili is required for growth on
Fe(III) oxides. These “microbial nanowires” are not required
for attachment to the insoluble electron acceptor; rather, they
function as electronic conduits to transfer electrons to the
Fe(III) oxides, extending the electron transfer capabilities of
the cell well beyond the outer surface (14). Pilus “nanowires”
also serve as electric conduits to mediate long-range electron
transfer across multilayer biofilms formed on anode elec-
trodes, which is required to maximize current production per
unit of anode surface area (15).
Biofilms on Fe(III) oxide. When G. sulfurreducens (2) was
grown under strictly anaerobic conditions at 30°C in freshwater
medium (7) with acetate (15 mM) as an electron donor and
with Fe(III) oxide coatings [prepared on borosilicate coverslips
(17) and providing 4.3 ? 0.7 ?mol of Fe(III) per coverslip
(mean ? standard deviation; n ? 3)] as the sole electron
acceptor, a biofilm grew on the Fe(III) coating (measured with
a crystal violet assay ), but planktonic growth was not
supported (Fig. 1A). Viability staining with a BacLight viability
kit (Molecular Probes) and confocal scanning laser microscopy
(CSLM) analyses (14) of 48-h biofilms revealed a structured
biofilm composed of cell clusters approximately 18 ? 1 ?m
high (Fig. 1B). Control coverslips without the Fe(III) oxide
coatings did not support biofilm growth (Fig. 1C), suggesting
that biofilm growth was not supported by any nutrient carried
over in the inoculum. Viability staining suggested that even
cells at a substantial distance from the Fe(III) oxide surface
remained metabolically active (Fig. 1B). This may be attrib-
uted to long-range electron transfer via the electrically con-
ductive pili, as previously proposed for long-range transfer to
the anode surface of microbial fuel cells (15). In contrast, a
previously described (14) mutant in which the gene coding for
PilA, the pilin structural subunit, was deleted grew poorly on
the Fe(III) oxide coatings (Fig. 1D) and produced 10-fold less
biomass than the wild type produced after 72 h (Fig. 2), and
complementation of the mutation in trans (14) restored the
biofilm phenotype (data not shown). These findings are con-
sistent with the previous finding that pili are required for
growth on Fe(III) oxide (14).
Biofilm formation when electron transfer to the Fe(III) ox-
ide surface is not required. Even though pili are not required
for growth with fumarate as an electron acceptor (14), addition
of fumarate (40 mM) to cultures with Fe(III) oxide-coated
coverslips, while having little impact on the biofilm biomass of
the wild type, increased the mutant biofilm biomass to approx-
imately one-half of the wild-type biomass (Fig. 2). More wild-
type biomass accumulated on glass coverslips when fumarate
was provided as the electron acceptor, but the biomass of the
pilin-deficient mutant biofilms remained approximately one-
half that of the wild-type biofilms (Fig. 2). These results dem-
onstrated that pili are required for optimal biofilm develop-
ment even when the surface is not the electron acceptor.
This conclusion was consistent with CLSM images of the
fumarate-grown biofilms (Fig. 3). In the presence of fumarate,
wild-type cells formed pillars that were 19 ? 1.5 and 22 ? 0.5
?m high on Fe(III) oxide-coated surfaces and glass surfaces,
respectively. In contrast, the maximum biofilm heights for the
pilin-deficient mutant were 7.6 ? 1.5 and 8.5 ? 1.4 ?m on the
same surfaces. This difference was not apparent in the first
24 h, when both the mutant and wild-type biofilms formed
short microcolonies. However, the wild-type microcolonies
continued to grow, and both the height and width of the col-
onies increased to form mature biofilms (Fig. 3). Viability
staining indicated that cells in all the layers of the wild-type
and mutant biofilms were alive, suggesting that fumarate dif-
* Corresponding author. Present address: Department of Microbi-
ology and Molecular Genetics, 2215 Biomedical Physical Sciences,
Michigan State University, East Lansing, MI 48824-4320. Phone: (517)
355-6463. Fax: (517) 353-8957. E-mail: firstname.lastname@example.org.
† Present address: Infectious Diseases Department, The Mount
Sinai School of Medicine, New York, NY 10029.
‡ Present address: Center for Adaptation Genetics and Drug Resis-
tance, Department of Molecular Biology and Microbiology, Tufts Uni-
versity School of Medicine, Boston, MA 02111.
?Published ahead of print on 8 December 2006.
fusion across the biofilms was not a limiting factor, as previ-
ously reported for other bacterial biofilms (16). These results
indicate that the pili of G. sulfurreducens play a role in the
development of the highly structured biofilms of G. sulfurre-
ducens that is unlikely to be related to the electrical conduc-
tivity of the pili.
Geobacter pili promote autoagglutination. Pili in various
bacteria mediate twitching motility during biofilm formation
(12). Other pili, such as the toxin-coregulated pili (TCP) of
Vibrio cholerae (13), and also the pili of G. sulfurreducens (14)
do not appear to be involved in motility. Rather, TCP are a
structural biofilm component that mediate cell interactions
leading to microcolony development during colonization of the
human intestine (3) or during biofilm formation on chitin sur-
faces (13). The ability of TCP to promote bacterial interactions
also enables TCP-expressing cells to autoagglutinate in vitro
(3). Similar agglutination studies were carried out with G.
sulfurreducens by growing cells with fumarate as the electron
acceptor at 25°C to induce pilus formation (14). The degree of
agglutination was assayed by measuring the optical density at
600 nm of the cells that remained in suspension and subtract-
ing the value obtained from the optical density of the culture
after disruption of the aggregates with agitation. After 72 h of
growth the wild-type strain formed large aggregates that set-
tled at the bottom of the culture vessel (Fig. 4). There was no
autoagglutination at 30°C, a temperature at which planktonic
cells do not express pili (14). The mutant in which pilA was
deleted did not agglutinate at 25°C (Fig. 4). Complementation
of the mutation with a wild-type copy of the pilA gene ex-
pressed in trans produced a strain that agglutinated at levels
FIG. 1. Biofilm formation on Fe(III) oxide coatings by G. sulfurreducens. (A) When provided as a sole electron acceptor for growth, the Fe(III)
oxide coating supported the growth of a biofilm, whose biomass increased steadily during the first 72 h, but did not support planktonic growth.
OD600, optical density at 600 nm. (B to D) Top view (at a 15oangle) (top panels) and side view (bottom panels) projections generated by CSLM
of 48-h wild-type (B) or pilin-deficient mutant (D) biofilms formed on an Fe(III) oxide-coated surface or of a wild-type biofilm on a control
coverslip without the Fe(III) oxide coating (C). Green indicates live cells, and red indicates dead cells. Yellow regions are areas where the two dyes
overlap. The substratum (coverslip) is at the bottom. Bars, 20 ?m.
FIG. 2. Average biomasses of mature wild-type (WT) and pilin-
deficient mutant (PilA?) biofilms formed on Fe(III) oxide coatings
[Fe(III)] or glass surfaces. Where indicated, the soluble electron ac-
ceptor fumarate also was present in the growth medium. The results
are averages for triplicate samples from two independent experiments.
OD600, optical density at 600 nm.
that were much higher than the levels observed for the wild- Download full-text
type strain (Fig. 4), consistent with the fact that genetic
complementation leads to overproduction of pili (14). These
results suggest that the pili of G. sulfurreducens participate in
cell-cell aggregation necessary for the development of micro-
colonies during biofilm differentiation.
Implications. The results presented here demonstrate that
in addition to serving as electric conduits for electron transfer
to Fe(III) oxides (14) and long-range electron transfer across
anode biofilms in G. sulfurreducens fuel cells (15), the G. sul-
furreducens pili also are required for maximum biofilm growth
even when electron transfer to an electron-accepting surface is
not required. This is an important consideration because the
overall rate of electron transfer to an electron-accepting sur-
face is dependent upon the number of metabolically active
cells that can stack on the surface. Thus, high rates of electron
transfer to an electron-accepting surface require not only the
electronic capabilities of the pili but also their structural at-
tributes that permit cells to stack at high densities on a given
surface. These considerations make it clear that further eval-
uation of the contributions of pili and other outer cell compo-
nents to the biofilm structure is essential in order to better
understand, and perhaps optimize, electron transfer to elec-
This research was supported by grants DE-FG02-02ER63423 and
DE-FC02-02ER63446 from the Office of Science (BER), U.S. Depart-
ment of Energy, and by award N00014-03-1-0405 from the Office of
Naval Research. G.R. acknowledges support provided by a postdoc-
toral fellowship from the Ministerio de Educacio ´n y Ciencia of Spain
and by the European Social Fund.
1. Bond, D. R., and D. R. Lovley. 2003. Electricity production by Geobacter
sulfurreducens attached to electrodes. Appl. Environ. Microbiol. 69:1548–
2. Caccavo, F., Jr., D. J. Lonergan, D. R. Lovley, M. Davis, J. F. Stolz, and M. J.
McInerney. 1994. Geobacter sulfurreducens sp. nov., a hydrogen- and acetate-
oxidizing dissimilatory metal-reducing microorganism. Appl. Environ. Mi-
3. Kirn, T. J., M. J. Lafferty, C. M. Sandoe, and R. K. Taylor. 2000. Delineation
of pilin domains required for bacterial association into microcolonies and
intestinal colonization by Vibrio cholerae. Mol. Microbiol. 35:896–910.
4. Lies, D. P., M. E. Hernandez, A. Kappler, R. E. Mielke, J. A. Gralnick, and
D. K. Newman. 2005. Shewanella oneidensis MR-1 uses overlapping pathways
for iron reduction at a distance and by direct contact under conditions
relevant for biofilms. Appl. Environ. Microbiol. 71:4414–4426.
5. Lovley, D. R. 2006. Bug juice: harvesting electricity with microorganisms.
Nat. Rev. Microbiol. 4:497–508.
6. Lovley, D. R., D. E. Holmes, and K. P. Nevin. 2004. Dissimilatory Fe(III) and
Mn(IV) reduction. Adv. Microb. Physiol.49:219–286.
7. Lovley, D. R., and E. J. P. Phillips. 1988. Novel mode of microbial energy
metabolism: organic carbon oxidation coupled to dissimilatory reduction of
iron or manganese. Appl. Environ. Microbiol. 54:1472–1480.
8. Nevin, K. P., and D. R. Lovley. 2000. Lack of production of electron-shuttling
compounds or solubilization of Fe(III) during reduction of insoluble Fe(III)
oxide by Geobacter metallireducens. Appl. Environ. Microbiol. 66:2248–2251.
9. Nevin, K. P., and D. R. Lovley. 2002. Mechanisms for accessing insoluble
Fe(III) oxide during dissimilatory Fe(III) reduction by Geothrix fermentans.
Appl. Environ. Microbiol. 68:2294–2299.
10. Nevin, K. P., and D. R. Lovley. 2002. Mechanisms for Fe(III) oxide reduction
in sedimentary environments. Geomicrobiol. J. 19:141–159.
11. Newman, D. K., and R. Kolter. 2000. A role for excreted quinones in extra-
cellular electron transfer. Nature 405:94–97.
12. O’Toole, G. A., and R. Kolter. 1998. Flagellar and twitching motility are
necessary for Pseudomonas aeruginosa biofilm development. Mol. Microbiol.
13. Reguera, G., and R. Kolter. 2005. Virulence and the environment: a novel
role for Vibrio cholerae toxin-coregulated pili in biofilm formation on chitin.
J. Bacteriol. 187:3551–3555.
14. Reguera, G., K. D. McCarthy, T. Mehta, J. S. Nicoll, M. T. Tuominen, and
D. R. Lovley. 2005. Extracellular electron transfer via microbial nanowires.
15. Reguera, G., K. P. Nevin, J. S. Nicoll, S. F. Covalla, T. L. Woodard, and D. R.
Lovley. 2006. Biofilm and nanowire production lead to increased current in
microbial fuel cells. Appl. Environ. Microbiol. 72:7345–7348.
16. Stewart, P. S. 2003. Diffusion in biofilms. J. Bacteriol. 185:1485–1491.
17. van Schie, P. M., and M. Fletcher. 1999. Adhesion of biodegradative anaer-
obic bacteria to solid surfaces. Appl. Environ. Microbiol. 65:5082–5088.
FIG. 4. Autoagglutination phenotypes of a wild-type strain (WT), a
pilus-deficient mutant (PilA?), and a genetically complemented mu-
tant (pRG5-pilA) grown under pilus-inducing conditions (25°C). The
results are the averages for triplicate samples from two independent
experiments. A600, absorbance at 600 nm.
FIG. 3. CSLM analyses of wild-type (A and C) and PilA?(B and
D) biofilms formed on Fe(III) oxide coatings (A and B) or glass
coverslips (C and D) in medium with fumarate. Green indicates live
cells, and red indicates dead cells. Yellow indicates dye overlap. The
images are three-dimensional top views (top panels) and side views
(bottom panels) reconstructed from the fluorescence patterns of the
series of two-dimensional optical sections collected by CSLM. Bars,
VOL. 189, 2007NOTES2127