AEM Accepts, published online ahead of print on 15 August 2014 Appl. Environ. Microbiol. doi:10.1128/AEM.02289-14 Copyright © 2014, American Society for Microbiology. All Rights Reserved.
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Enhanced uranium immobilization and reduction by Geobacter sulfurreducens biofilms
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Dena L. Cologgi,a* Allison M. Speers,a Blair A. Bullard,a Shelly D. Kelly,b and Gemma Regueraa#
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Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing,
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Michigan a; EXAFS Analysis, Bolingbrook, Illinois b
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Running Head: Uranium reduction by Geobacter biofilms
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Keywords: metal reduction, uranium, biofilms, pili, c-cytochromes, bioremediation
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#
Address correspondence to Gemma Reguera, [email protected]
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* Present address: Dena Cologgi, department of Civil and Environmental Engineering,
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University of Alberta, Edmonton, Alberta, Canada.
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ABSTRACT
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Biofilms formed by dissimilatory metal reducers are of interest to develop permeable biobarriers
20
for the immobilization of soluble contaminants such as uranium. Here we show that biofilms of
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the model uranium-reducing bacterium Geobacter sulfurreducens immobilized substantially
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more U(VI) than planktonic cells and for longer periods of time, reductively precipitating it to a
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mononuclear U(IV) phase involving carbon ligands. The biofilms also tolerated high and
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otherwise toxic concentrations (up to 5 mM) of uranium, consistent with a respiratory strategy
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that also protected the cells from uranium toxicity. The enhanced ability of the biofilms to
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immobilize uranium correlated only partially with the biofilm biomass and thickness and
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depended greatly on the area of the biofilm exposed to the soluble contaminant. By contrast,
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uranium reduction depended on the expression of Geobacter conductive pili and, to a lesser
29
extent, to the presence of the c-cytochrome OmcZ in the biofilm matrix. The results support a
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model in which the electroactive biofilm matrix immobilizes and reduces the uranium in the top
31
stratum. This mechanism prevents the permeation and mineralization of uranium in the cell
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envelope, thereby preserving essential cellular functions and enhancing the catalytic capacity of
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Geobacter cells to reduce uranium. Hence, the biofilms provide cells with a physical and
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chemical protective environment for the sustained immobilization and reduction of uranium that
35
is of interest for the development of improved strategies for the in situ bioremediation of
36
environments impacted by uranium contamination.
37 38
INTRODUCTION
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In their natural environment microorganisms are most often found as surface-attached
40
communities or biofilms (1, 2). Biofilm cells are encased in a matrix of exopolymeric substances
41
(EPS) such as polysaccharides, nucleic acids, lipids, and proteins, which promote adhesion to
42
surfaces and mechanically stabilize the multilayered communities (3). In addition, the EPS
2
43
matrix functions as a hydrated, catalytic microenvironment for the cells that promotes the
44
adsorption of nutrients and, in some cases, their extracellular processing to facilitate their
45
assimilation (3). Although the biofilm structure is dynamic and can change to minimize mass
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transfer limitations (4), gradients of nutrients and waste products may form within the biofilms
47
and, as a result, the chemical composition of the matrix, and the physiology of the biofilm cells,
48
is heterogeneous (5). This unique physical and chemical microenvironment also makes the
49
biofilm cell physiology substantially different from that of planktonic cells and leads to some
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general, biofilm-specific traits such as increased resistance to antimicrobials (6), greater
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catalytic rates (7), and increased metabolic productivity (8).
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Multimetal resistance is also a common biofilm trait (9) and, for this reason, biofilms have
53
attracted interest for applications in metal bioremediation (10). The polyionic nature of the
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biofilm matrix can provide, for example, 20- to 30-fold more charged groups for metal sorption
55
than planktonic cells (11). This limits the diffusion of the metals within the biofilm and their
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permeation inside the cells, thus reducing the susceptibility of the biofilm cells to metal toxicity
57
compared to their planktonic counterparts (9). The chemical and physiological heterogeneity of
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the biofilms also establishes pH and redox gradients across the biofilm matrix, which may
59
influence metal speciation (9). The chemistry of some metals is such that their speciation affects
60
their solubility and, therefore, their potential for spread and their bioavailability. This is
61
particularly important for uranium (U), which often persists in contaminated groundwater and
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sediments as the soluble uranyl cation (UO22+) containing the oxidized U(VI) species. Its
63
solubility facilitates the spread of the contaminant plume far away from the source (12) and
64
results in volumes of contaminated groundwater and sediments too large to permit standard
65
‘excavation and removal’ and ‘pump and treat’ remediation approaches (13). Hence, a
66
promising bioremediation approach would be to immobilize the U(VI) contaminant and reduce it
67
to the U(IV) species, which is less soluble and can precipitate out of solution under the right
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redox conditions and pH (12). This prevents the migration of the contaminant, reduces its
3
69
bioavailability, and minimizes the potential for exposure to humans and other living components
70
of the ecosystem.
71
One way to reduce U(VI) to U(IV) is by the action of some dissimilatory metal-reducing
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bacteria, which can gain energy for growth by coupling the oxidation of various electron donors
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to the reduction of the uranyl cation (14-16). This metabolic ability can be stimulated in situ with
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field-scale additions of electron donors and results in the removal of the uranyl cation from the
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groundwater and its accumulation in the sediments as sparingly soluble, less mobile U(IV)
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minerals (17-21). Field-scale additions of electron donors often stimulate the growth and activity
77
of dissimilarity metal-reducing microorganisms within the family Geobacteraceae (17, 19, 20,
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22). Studies with the model representative Geobacter sulfurreducens indicate that the reduction
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of U by these organisms is extracellular; this prevents the permeation and reductive
80
precipitation of the radionuclide inside the cell envelope, thereby protecting the cell from its
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toxicity (23). The extracellular reduction of U by G. sulfurreducens is primarily mediated by hair-
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like protein filaments or pili and, to a lesser extent, by c-cytochromes of the outer membrane
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(23). The pili of Geobacter are conductive (24) and their expression levels are linearly
84
proportional to the amount of U reductively precipitated by planktonic cells (23). Pili expression
85
in G. sulfurreducens is also required to form an electroactive biofilm matrix (25, 26), where the
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cells are encased in a exopolysaccharide matrix containing c-cytochromes (27). This suggests
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that Geobacter biofilms may be particularly suitable to catalyze the speciation of U while
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providing a microenvironment that also protects cells from its toxicity.
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Biofilms are particularly prevalent at the matrix-well screen interface and within rock fractures
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in subsurface environments contaminated with U, thereby influencing its mobility and speciation
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(13). However, the contribution of Geobacter biofilms to these processes and their potential use
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as bioremediation tools has not been fully investigated. This contrasts with the availability of
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studies about U transformations mediated by Geobacter planktonic cells (14, 15, 23, 28, 29) or
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by biofilms formed by other metal-reducing bacteria such as Desulfovibrio (30, 31) and
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Shewanella (32, 33). Hence, we investigated U transformations mediated by biofilms of G.
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sulfurreducens. The results indicate that the biofilms have enhanced capacity to immobilize and
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reduce U compared to planktonic cells and also tolerate exposure to higher concentrations of
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the contaminant for prolonged periods of time. Furthermore, microscopic, genetic, and
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spectroscopic studies of the U-reducing biofilms indicated that the radionuclide was reduced
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extracellularly in a catalytic process influenced by the biofilm structure and the presence of
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redox components of the biofilm matrix, most significantly the conductive pili. These findings
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support the notion that Geobacter biofilms contribute to the immobilization and reduction of U in
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the subsurface and highlight their potential use as permeable biobarriers for the in situ
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bioremediation of U.
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MATERIALS AND METHODS
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Strains and culture conditions. Wild-type (WT) Geobacter sulfurreducens PCA (ATCC
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51573), a pilin-deficient mutant (24) (herein designated pilA), and its genetically complemented
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strain pRG5::pilA (24) (herein designated pilA+) were routinely cultured in fresh water (FW)
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medium (34) modified as previously described (23) and supplemented with 15 mM acetate and
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40 mM fumarate (FWAF). The medium was dispensed into tubes or serum bottles, sparged with
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N2:CO2 (80:20), sealed with butyl rubber stoppers (Bellco) and aluminum tear-off seals
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(Wheaton), and autoclaved 30 minutes. Unless otherwise indicated, incubations were at 30 oC.
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Biofilm cultures. Biofilms were either grown on 6-well, cell-culture-treated polystyrene
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plates (Corning) or on glass coverslips assembled vertically, four at a time, onto rubber
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stoppers, as described previously (26). Briefly, the glass coverslips used for the stopper
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assemblies were first acid-washed overnight in a bath of 50/50 (v/v) HCl/NO3- or 15% (v/v)
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HCl/H2O and rinsed thoroughly with ddH2O before inserting them into slits cut on inverted
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stoppers. Each coverslip assembly was placed upside down (coverslips down) in a sterile 50 ml
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conical tube (Corning) and submerged in FW medium lacking vitamins, minerals, acetate, and
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fumarate before sterilizing it by autoclaving, as previously described (26). The assembly was
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then transferred to a clean conical tube. The biofilm chambers (either wells of plates or conical
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tubes with coverslip assemblies) were filled with 6 or 20 ml of FWAF medium, respectively, and
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inoculated to a final OD600 of 0.04 with an early stationary-phase FWAF culture. The chambers
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were incubated at 30 °C for 24, 48, or 72 h, as specified, inside a vinyl glove bag (Coy Labs)
126
with a H2:CO2:N2 (7:10:83) atmosphere. At the end of the incubation period, the culture
127
supernatant fluids were decanted from the biofilm vessels and the biofilms were washed once
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with sterile, anaerobically-prepared wash buffer (29).
129
When indicated, the biofilm biomass in the samples was estimated as total cell protein.
130
To do this, the biofilm biomass was scraped off from the surface with a sterile plastic spatula,
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harvested by centrifugation (5 min, 12,000 x g), and the biofilm cells were then lysed in 2M
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NaOH at 100 oC for 1 h. The lysate solution was allowed to cool before neutralizing it with an
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equal volume of 2M HCl. After another cycle of centrifugation to remove cellular debris, the
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supernatant, which contained the soluble protein released after lysing the cells, was analyzed
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for protein content using a Pierce Microplate BCA Protein Assay Kit (reducing reagent
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compatible, Thermo Scientific) with BSA standards, according to the manufacturer’s
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specifications. Protein was measured as an OD562 on a Tecan Sunrise Plate Reader (Tecan,
138
Inc.).
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U immobilization assays with resting planktonic cell suspensions and biofilms.
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The ability of planktonic or biofilm cells to immobilize U was assayed by monitoring the removal
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of U(VI), provided as uranyl acetate (1 mM, or as indicated), at 30 oC by resting planktonic cells
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and biofilms, using protocols adapted from those described previously (23, 29). Planktonic cells
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were harvested from mid-exponential cultures (OD600, 0.3-0.5) grown for 48 h under pili-inducing
145
conditions (25 oC) and suspended in 10 ml of reaction buffer (OD600, 0.1) as described before
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(23). Resting biofilms were prepared by first growing the surface-attached communities in
6
147
FWAF medium for 24, 48, or 72 h on the coverslip-stopper assemblies described above. The
148
culture broth of the biofilm culture was then decanted gently and the coverslip assembly was
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rinsed gently with sterile, anaerobically-prepared wash buffer (29). To initiate the assay, the
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assemblies were submerged upside down in 50-ml conical tubes containing 20 ml of reaction
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buffer (23, 29) supplemented with 20 mM sodium acetate and 1 mM uranyl acetate (Electron
152
Microscopy Sciences), which had been prepared from stock solutions in 30 mM bicarbonate
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buffer. Heat-killed planktonic cells (23) and uninoculated biofilm controls were also included to
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rule out any abiotic removal activity and/or absorption. Resting planktonic cells, resting biofilms,
155
and the controls were incubated for up to 24 h at 30 °C. Supernatant samples (500 l) were
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periodically removed during incubation, filtered (0.22 µm Millex-GS filter, Millipore), acidified in
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2% nitric acid (500 l), and stored at -20 °C. All procedures were performed inside an anaerobic
158
glove bag, as described above. The concentration of U(VI) in the acidified samples was
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measured using a Platform Inductively Coupled Plasma Mass Spectrometer (ICP-MS)
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(Micromass,Thermo Scientific) or a Kinetic Phosphorescence Analyzer (KPA) (Chemchek) and
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provided a measure of the total U immobilized in each sample. For comparisons, the amount of
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U immobilized by the planktonic and biofilm cells was also normalized by the cell biomass,
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which was calculated as the amount of total cell protein released from the samples after lysis
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with 2M NaOH (100 oC for 1 h) and neutralization with 2M HCl, as described above.
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U valence and speciation by X-ray Absorption Spectroscopy (XAS) analyses. The
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valence and speciation of the U immobilized by resting biofilms was estimated by XANES (X-ray
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Absorption Near Edge Spectroscopy) and EXAFS (Extended X-ray Absorption Fine Structure
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Analysis), respectively. For these analyses, biofilms grown on coverslip-stopper assemblies and
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exposed to U for 24 h, as described above, were rinsed gently with wash buffer, scraped off the
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coverslips, and suspended in 2 ml of reaction buffer. The biofilm biomass was then harvested
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by centrifugation (12,000 x g, 10 min), loaded into custom-made plastic holders, and stored at -
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80 °C (23). All procedures were carried out in an anaerobic chamber and samples were kept
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frozen during XAS measurements. The XAS measurements were performed with a
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multielement Ge detector in fluorescence mode using the PNC-CAT beamline 20-BM at the
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Advanced Photon Source (Argonne National Laboratory) and standard beamline parameters, as
177
described elsewhere (35). XANES measurements were used to calculate the fraction of the
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immobilized U that had been reduced to U(IV) by linear combination fitting of the spectrum with
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U(VI) and U(IV) standards. The spectra were energy aligned using a simultaneously measured
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uranyl nitrate standard. The U LIII-edge EXAFS spectrum was also collected for the biofilm
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samples and modeled to determine the atomic coordination about U, as previously described
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(23). The number of axial oxygen (Oax) atoms calculated in the EXAFS spectral analyses was
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also used to estimate the amount of U(VI) and U(IV) in the samples, as there are two double-
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bonded Oax atoms for each U(VI) atom in the uranyl cation (Oax=U=Oax) and none for U(IV)
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(36).
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Vitality fluorescent assays. The respiratory activity of 48-h old biofilms after exposure
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to various concentrations (1, 2.5, and 5 mM) of uranyl acetate for 24 h was assayed using the
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fluorescent RedoxSensor vitality kit (Invitrogen) in reference to biofilms not exposed to uranyl
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acetate (0 mM controls), with protocols adapted from those described previously (23). The
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biofilms were first grown for 48 h on coverslip assemblies, rinsed gently with reaction buffer, and
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incubated in reaction buffer in the presence or absence of the uranyl acetate. The experiment
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also included negative controls (0* mM) in which the 0 mM biofilms were first treated for 5 min
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with electron transport chain uncouplers (a mix of sodium azide [10 mM] and carbonyl cyanide
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3-chlorophenylhydrazone or CCCP [10 μM]) provided in RedoxSensor vitality kit (Invitrogen).
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After 24 h of incubation, the reaction buffer was decanted from the tubes and the assemblies
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were washed once with wash buffer. The biofilm biomass was scraped from the coverslips and
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suspended in 1 ml of reaction buffer, vortexed briefly, mixed 1:1 with the Redox dye solution,
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and incubated at room temperature for 10 min before measuring the dye’s fluorescence (490
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nm excitation, 520 nm emission) using a SpectraMax M5 plate reader (Molecular Devices).
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Separate aliquots of the biofilm samples were stained with SYTO 9 (Invitrogen) following
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manufacturer’s recommendations to determine the total biofilm biomass. The respiratory activity
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of the biofilms was estimated as the vitality index, i.e., the relative fluorescence emission from
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the RedoxSensor dye normalized by the fluorescence emission from the Syto9 dye for 6
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replicate samples (3 biological samples and two technical replicates for each). Statistically
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significant (p < 0.05, one star; p < 0.005, two stars) changes in vitality indexes were determined
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in pairwise comparisons to the 0 mM control biofilms using the t-test function of the Microsoft
208
Excel software.
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Microscopy. An Scanning Electron Microscope (SEM) was used to image 48-h old WT
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biofilms, which were grown on coverslip assemblies as described above except that round (12-
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mm diameter), rather than square, glass coverslips were used. The biofilms were incubated at
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30 oC in reaction buffer in the presence or absence of 1 mM uranyl acetate for 24 h and washed
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once in wash buffer before fixing the cells at 4°C for 1-2 h in 4% glutaraldehyde. After this, the
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biofilms were rinsed briefly in 0.1 M sodium phosphate buffer and then dehydrated in a series of
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ethanol washes (25%, 50%, 75%, 95%, 10 minutes each) followed by three 10-min washes in
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100% ethanol. The samples were critical point dried using a Blazers 010 critical point dryer
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(Blazers Union Ltd.) with liquid CO2 as the transitional fluid. Once dry, the coverslips containing
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the biofilm samples were mounted on aluminum stubs using epoxy glue and coated with ~10
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nm of osmium using a NEOC-AT osmium coater (Meiwafosis Co., Ltd.). Samples were imaged
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with a JEOL JSM-7500F SEM equipped with an Energy Dispersive Spectroscopy (EDS) 30mm2
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detector crystal, which was used for elemental analyses.
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Confocal Laser Scanning Microscopy (CLSM) was also used to examine the biofilms
224
and generate 3D-images for structural analyses. The WT biofilms were grown for 24, 48, and 72
225
h on 6-well plates and the pilA and pilA+ biofilms were grown for 48 h. At the end of the
226
incubation period, the culture supernatant fluid from each well was carefully decanted, and the
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biofilms were stained for 15 min with the LIVE/DEAD® BacLight™ Bacterial Viability Kit
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(Invitrogen) dye solution, following the manufacturer’s recommendations. After staining, the
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biofilms were washed once in phosphate buffer saline (PBS) and imaged on a Zeiss Pascal
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LSM microscope (Carl Zeiss Microscopy, LLC) equipped with an Achroplan 40x/0.80W
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submersible objective. Images were collected every 1.14 µm, and side and top projections of
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the biofilms were created using the Zeiss LSM Image Browser software (Carl Zeiss Microscopy,
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LLC). Approximately 6-10 distinct fields of view (1,024 x 1,024 pixels, 0.22 µm/pixel) were
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imaged for each of three independent biofilm replicates and the micrographs were analyzed with
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the COMSTAT image analysis software using connected volume filtration to remove noise in the
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data, as described previously (4). The biofilm structural parameters quantified with the
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COMSTAT analyses are listed in Table S2.
238 239
SDS-PAGE and heme-stain of proteins of the biofilm EPS matrix. The
240
exopolysaccharide matrix (EPS) of 48-h biofilms grown on 6-well plates was extracted using a
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modification of previously described protocols (27, 37). Briefly, biofilms were scraped off the
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wells, collected in reaction buffer, and the solution was centrifuged for 10 min at 13,000 x g.
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After decanting the supernatant fluid, the biofilm pellet was suspended in 1/5 volume of TNE (10
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mM Tris-HCl [pH 7.5], 100 mM NaCl, 5 mM EDTA) and vortexed for 1 min. SDS was then
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added to a final concentration of 0.1% (w/v), and the solution was mixed at room temperature
246
for 5 min. The samples were then passed 10 times through an 18G needle, and centrifuged at
10
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15,500 x g for 20 min to collect the insoluble sheared biological fraction. The resulting pellet,
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which predominantly contained the biofilm matrix and associated proteins, was washed 5 times
249
to remove any SDS before suspending it in 10 mM Tris-HCl buffer (pH 7.5).
250
Heme-containing proteins in the EPS biofilm matrix were also analyzed, as previously
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described (27). The protein content in the matrix sample was determined with the bicinchoninic
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acid (BCA) protein assay, as described above, and the equivalent of 20 µg of protein of EPS
253
sample was boiled in SDS sample buffer for 10 min before electrophoretic analysis in a 12%
254
Mini-Protean TGX gel (BioRad) at 250 V for 30 min. The Novex Sharp markers (Invitrogen)
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were used as molecular weight standards. Heme-containing proteins were visualized on the gel
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after N,N,N’,N’-tetramethylbenzidine staining, as described previously (23, 38). A duplicate gel
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was run in parallel and stained for total protein using Coomassie Brilliant Blue G-250 (BioRad)
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according to the manufacturer’s recommendations.
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RESULTS AND DISCUSSION
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Enhanced U immobilization and tolerance by biofilms. The kinetics of U
262
immobilization were investigated by measuring the amount of U, provided as uranyl acetate,
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removed from solution by resting 48-h old biofilms and in reference to pili-expressing planktonic
264
cells (Fig. 1A). Heat-killed planktonic cell controls and uninoculated biofilm vessel controls were
265
also included to rule out any abiotic precipitation of U either due to components of the medium
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(such as phosphate) or adsorption to the biofilm coverslip assemblies and vessels used for the
267
assays (Fig. 1A). The kinetics of U removal by the biofilms was linear throughout the 24-h long
268
assay (~10 µM/h; R² = 0.967). By contrast, the planktonic cells stopped removing U from
269
solution after 12 h and all the U immobilized by the planktonic cell biomass was solubilized
270
again after 24 h of incubation. The enhanced capacity of the biofilms to immobilize U compared
271
to planktonic cells (Fig. 1A) cannot be explained by differences in cell biomass because the
272
rates of U immobilization per h, once normalized by the total cell protein, were 6-times higher in
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the biofilms (~2.8 µmol U immobilized per mg protein per h) than in planktonic cells (~0.45
274
µmol/mg/h). Furthermore, the biofilms sustained the rates of U immobilization 2 times longer (24
275
h) than their planktonic counterparts. As a result, the yields of U immobilization by the biofilms at
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the end of the 24-h assay were 12-times greater (~66 µmol of U immobilized per mg of biofilm
277
biomass protein) than the maximum yields measured in the planktonic cells after 12 h (~5.4
278
µmol of U per mg of protein).
279
The sustained immobilization of U by 48-h old biofilms for 24 h suggests that the biofilm
280
cells remained viable and metabolically active throughout the assay. To investigate this, we
281
used the fluorogenic RedoxSensorTM dye to measure the respiratory activities of the biofilms
282
after exposure to increasing concentrations of U (1, 2.5, and 5 mM) for 24 h and in reference to
283
biofilms incubated similarly but without uranium (0 mM) (Fig 1B). Interestingly, the vitality index
284
of the biofilms exposed to 1 mM U was greater than the unexposed 0 mM controls.
285
Furthermore, increasing the concentrations of U (2.5 and 5 mM) did not decrease the
286
respiratory activities of the biofilms to or below unexposed biofilm levels. The measured vitality
287
correlated well with the respiratory activity of the biofilms because pretreating the biofilms with
288
sodium azide and CCCP (0* mM controls), which are chemicals that uncouple the electron
289
transport chain, reduced the vitality by 80%. The dye used in these experiments yields green
290
fluorescence when modified by the bacterial reductases, which are mostly located in the
291
electron transport system of the cell envelope (39, 40). It is unlikely that the increases in the
292
biofilm’s respiratory rate with U were due to a cellular response of the cells to U toxicity,
293
because the vitality index also serves as a proxy of viability and overall cell health in G.
294
sulfurreducens cells after U exposure (23). For example, exposure of pili-expressing planktonic
295
cells to 1 mM U for only 6 h reduces the respiratory activity of the cells by 70%, because of the
296
toxic effects of the U that permeates and mineralizes in the cell envelope (23). Yet when the
297
cells are encased in the protective biofilm microenvironment they tolerated high, otherwise toxic
298
levels of U, and their metabolic activities were stimulated as well.
12
299
U reduction by the biofilms. As the reaction of interest during U bioremediation is the
300
reduction of the soluble U(VI) species to the less mobile U(IV), we used XANES to investigate if
301
any of the U immobilized by the biofilms had been reduced to U(IV). For these experiments, the
302
48-h old biofilms studied in Fig. 1A were collected at the end of the 24-h U challenge and the
303
valence of the biofilm-associated U was analyzed by XANES in reference to U(VI) and U(IV)
304
standards. Approximately 65% (± 5%) of the U immobilized by the biofilms was in the reduced
305
state, U(IV) (Table S1). Consistent with the reductive precipitation of U by the biofilms, SEM-
306
EDS analyses revealed needle-like, extracellular precipitates coating the microcolonies (Fig. 2A)
307
that contained U (Fig. S1). The preferential localization of U minerals on the top regions of the
308
biofilm is consistent with the metabolic stratification of cells within electroactive biofilms of G.
309
sulfurreducens (41). As the biofilms grow, their thickness increases and a gradient of acetate is
310
formed across the biofilm. This concentrates the metabolically active cells in the top regions of
311
the biofilm, where acetate is more available. As a result of this metabolic stratification, U is
312
preferentially reduced in the metabolically-active, top biofilm stratum (41).
313
SEM micrographs also revealed the presence of extracellular filaments interspersed with
314
the U precipitates (Fig. 2B). The dense filament network was more clearly visible in biofilm
315
controls not exposed to U (Fig. 2C). Some filaments had diameters (< 4 nm) that closely
316
matched the diameters reported for the conductive pili of G. sulfurreducens (24), whereas
317
others had larger diameters (ca. 15-20 nm) within the ranges reported for dehydrated EPS
318
fibers (27). The conductive pili of G. sulfurreducens are the primary sites for extracellular
319
immobilization and reduction of U by pili-expressing planktonic cells (23) and are also required
320
for the formation of electroactive biofilms (25, 26). In addition to containing the conductive pili,
321
the EPS matrix of G. sulfurreducens also anchors several c-cytochromes involved in metal
322
reduction (27). Thus, the Geobacter matrix provides an extended matrix for U immobilization but
323
also contains redox-active components which could mediate its reduction.
13
324
U immobilization and reduction during biofilm development. As the biofilms
325
develop, their thickness increases and more biofilm matrix and matrix-associated redox
326
components are available to immobilize and reduce U, respectively. Hence, we hypothesized
327
that older and thicker biofilms would immobilize and reduce more U than younger and thinner
328
biofilms. For this reason, we studied U immobilization and reduction as a function of biofilm age
329
(24-h, 48-h, and 72-h old biofilms) (Table S1). The biofilms were also imaged by confocal
330
microscopy (Fig. 4) and the confocal micrographs were analyzed with the COMSTAT software
331
(4) to estimate the thickness and other structural parameters of the biofilms (Table S2).
332
We first studied U immobilization during biofilm development. Although the capacity of
333
the biofilms to immobilize U increased steadily as they aged (Fig. 3A) and their thickness
334
increased (Fig. 4), the amount of U immobilized per biomass unit did not change significantly
335
during biofilm development (Fig. S3A). For example, young (24-h old) biofilms had less biofilm
336
biomass, thickness, and surface area (cells per biofilm layer) than older (48-h and 72-h old)
337
biofilms (Fig. S4, panels A-C). Yet despite being less dense, they immobilized more U per
338
biofilm biomass unit than older biofilms (Fig. S3A). Interestingly, surface coverage by all of the
339
biofilms was similar and neared confluence (81-92%) (Fig. S4D). The surface to volume ratio
340
also remained relatively constant during biofilm development (Fig. S4E), indicating that the area
341
of the biofilm exposed to the liquid millieu relative to the biofilm volume is the same. This also
342
maintained the biofilm topography constant, as indicated by the similar roughness coefficients
343
measured in all the biofilms (Fig. S4F). Taken together, the results support a model in which
344
achieving biofilm confluence early on in biofilm development is critical to promote U
345
immobilization. As the biofilms age, they adapt their structure to maintain a topography that
346
maintains sufficient areas of the biofilms exposed to the liquid milleu and maintained similar
347
levels of U immobilization per biomass unit. This model also implies that U is preferentially
348
immobilized in the biofilm regions exposed to the liquid milley, which is further supported by the
14
349
preferential localization of U on the outer layers of the biofilms revealed in SEM micrographs
350
(Fig. 2).
351
The fraction of the immobilized U that was reduced to U(IV) was also estimated by
352
XANES (Table S1). Interestingly, the yields of U reduction increased linearly with biofilm age
353
(Fig. 3A), even when normalized by the biofilm biomass (Fig. S3A). None of the biofilm
354
structural parameters determined in the COMSTAT analyses fit the linear trend of U reduction
355
(Fig. S4). Parameters important for U immobilization (surface coverage, surface to volume
356
ration, and roughness coefficient) were, for example, unaffected by biofilm age. The biofilm
357
parameters that changed were those related to biofilm biomass (biofilm biomass, thickness, and
358
surface area) but they all stabilized in the 48-h and 72-h old biofilms rather that increasing
359
linearly. By contrast, the total protein content of the biofilms, which accounts for protein
360
contributed by cells and by proteinaceous components of the biofilm matrix, increased linearly
361
as the biofilms aged (Fig. S5). Biofilm formation is a developmental process, consisting of
362
specific stages such as attachment, microcolony formation, and biofilm maturation (42). As they
363
transition from one stage of biofilm development to the next, the biofilms increase their
364
thickness and biomass but also undergo extensive gene reprogramming so that specific biofilm
365
components and activities are expressed (9, 43). Pili expression in G. sulfurreducens is, for
366
example, required to transition from young, monolayered biofilms to thicker biofilms composed
367
of several layers of cells (26) and this, in turn, is required to maintain the electroactivity of the
368
biofilms (25). Hence, as the Geobacter biofilms age, they express more redox-active
369
components in the biofilm matrix and this, in turn, may increase their capacity to reduce the
370
immobilized U to U(IV).
371 372
Role of the redox components of the biofilm matrix in U immobilization and
373
reduction. As the conducive pili of G. sulfurreducens are the primary site for U immobilization
374
and reduction in planktonic cells (23) and also are required for biofilm formation (25, 26), we
15
375
compared the capacity of 48-h biofilms of the WT, a pilin-deficient mutant (pilA), and its
376
genetically complemented strain pilA+ to immobilize and reduce U (Fig. 3B). The pilA biofilms
377
had a diminished capacity to immobilize and reduce U (p = 0.07) compared to WT biofilms of
378
the same age. The defect was due to the pilus deficiency because complementing the mutation
379
in trans (pilA+) restored the phenotypes and promoted U immobilization and reduction to levels
380
comparable to the WT biofilms.
381
The defect in pili production in the pilA biofilms also prevented the biofilms from growing
382
in thickness (Fig. 4) and lead to ~ 50% decreases in other biomass-related structural
383
parameters such as biomass volume and surface area (cells per biofilm layer) compared the
384
WT biofilms (Table S2 and Fig. S4A-C). These defects are consistent with the reported role of
385
the pili at promoting cell-cell aggregation during biofilm formation and providing the structural
386
support required to build multilayered biofilms (26). However, despite the biomass and pili
387
defects, the pilA biofilms immobilized more U per biomass unit than the WT biofilms.
388
Furthermore, genetic complementation of the pilA mutation in the pilA+ strain resulted in denser
389
biofilms (Fig. 4), consistent with the increased ability of the pilA+ hyperpiliated strains to
390
aggregate (26). Yet despite the increased biofilm biomass, the pilA+ biofilms immobilized less U
391
per biofilm biomass unit than any other biofilm (Fig. S3B). The lack of a clear correlation
392
between biofilm biomass and U immobilization in the pilA, WT, and pilA+ biofilms may reflect
393
differences in diffusion rates of electron donors and acceptors, as more diffusional constraints
394
are expected in thicker (pilA+>WT) than in thinner (pilA) biofilms. Consistent with this, the pilA+
395
biofilms had lower surface to volume ratios than any other biofilms (Fig. S4E and Table S2),
396
indicating that less surface area of the biofilm volume is exposed to the liquid medium and
397
available for electron donors and acceptors diffusion. Furthermore, the roughness coefficient,
398
which provides a measure of the spatial heterogeneity of the biofilms as variations in biofilm
399
thickness, was also lowest in the pilA+ biofilms (Fig, S4F). This indicates that the very dense
400
pilA+ biofilms have a relative uniform topography (very low roughness coefficient, ~0.06),
16
401
whereas the thin pilA biofilms formed the most heterogenous biofilms of all (highest roughness
402
coefficient, ~0.32, almost twice the WT coefficients) (Table S2). Hence, the more heterogenous
403
structure of the pilA and WT biofilms also maximizes the biofilm surface area available for
404
diffusion of electron donors and acceptors.
405
Although the pili-deficient pilA biofilms immobilized more U per biomass unit than any
406
other strain, they reduced a lower fraction (~38%) of the immobilized U to U(IV) than the WT
407
(~63%) and pilA+ (~76%) biofilms per biomas unit (Fig. S3). The capacity of the biofilms to
408
reduce U (pilA
1
Enhanced uranium immobilization and reduction by Geobacter sulfurreducens biofilms
2 3
Dena L. Cologgi,a* Allison M. Speers,a Blair A. Bullard,a Shelly D. Kelly,b and Gemma Regueraa#
4 5
Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing,
6
Michigan a; EXAFS Analysis, Bolingbrook, Illinois b
7 8
Running Head: Uranium reduction by Geobacter biofilms
9 10
Keywords: metal reduction, uranium, biofilms, pili, c-cytochromes, bioremediation
11 12
#
Address correspondence to Gemma Reguera, [email protected]
13 14
* Present address: Dena Cologgi, department of Civil and Environmental Engineering,
15
University of Alberta, Edmonton, Alberta, Canada.
16 17
18
ABSTRACT
19
Biofilms formed by dissimilatory metal reducers are of interest to develop permeable biobarriers
20
for the immobilization of soluble contaminants such as uranium. Here we show that biofilms of
21
the model uranium-reducing bacterium Geobacter sulfurreducens immobilized substantially
22
more U(VI) than planktonic cells and for longer periods of time, reductively precipitating it to a
23
mononuclear U(IV) phase involving carbon ligands. The biofilms also tolerated high and
24
otherwise toxic concentrations (up to 5 mM) of uranium, consistent with a respiratory strategy
25
that also protected the cells from uranium toxicity. The enhanced ability of the biofilms to
26
immobilize uranium correlated only partially with the biofilm biomass and thickness and
27
depended greatly on the area of the biofilm exposed to the soluble contaminant. By contrast,
28
uranium reduction depended on the expression of Geobacter conductive pili and, to a lesser
29
extent, to the presence of the c-cytochrome OmcZ in the biofilm matrix. The results support a
30
model in which the electroactive biofilm matrix immobilizes and reduces the uranium in the top
31
stratum. This mechanism prevents the permeation and mineralization of uranium in the cell
32
envelope, thereby preserving essential cellular functions and enhancing the catalytic capacity of
33
Geobacter cells to reduce uranium. Hence, the biofilms provide cells with a physical and
34
chemical protective environment for the sustained immobilization and reduction of uranium that
35
is of interest for the development of improved strategies for the in situ bioremediation of
36
environments impacted by uranium contamination.
37 38
INTRODUCTION
39
In their natural environment microorganisms are most often found as surface-attached
40
communities or biofilms (1, 2). Biofilm cells are encased in a matrix of exopolymeric substances
41
(EPS) such as polysaccharides, nucleic acids, lipids, and proteins, which promote adhesion to
42
surfaces and mechanically stabilize the multilayered communities (3). In addition, the EPS
2
43
matrix functions as a hydrated, catalytic microenvironment for the cells that promotes the
44
adsorption of nutrients and, in some cases, their extracellular processing to facilitate their
45
assimilation (3). Although the biofilm structure is dynamic and can change to minimize mass
46
transfer limitations (4), gradients of nutrients and waste products may form within the biofilms
47
and, as a result, the chemical composition of the matrix, and the physiology of the biofilm cells,
48
is heterogeneous (5). This unique physical and chemical microenvironment also makes the
49
biofilm cell physiology substantially different from that of planktonic cells and leads to some
50
general, biofilm-specific traits such as increased resistance to antimicrobials (6), greater
51
catalytic rates (7), and increased metabolic productivity (8).
52
Multimetal resistance is also a common biofilm trait (9) and, for this reason, biofilms have
53
attracted interest for applications in metal bioremediation (10). The polyionic nature of the
54
biofilm matrix can provide, for example, 20- to 30-fold more charged groups for metal sorption
55
than planktonic cells (11). This limits the diffusion of the metals within the biofilm and their
56
permeation inside the cells, thus reducing the susceptibility of the biofilm cells to metal toxicity
57
compared to their planktonic counterparts (9). The chemical and physiological heterogeneity of
58
the biofilms also establishes pH and redox gradients across the biofilm matrix, which may
59
influence metal speciation (9). The chemistry of some metals is such that their speciation affects
60
their solubility and, therefore, their potential for spread and their bioavailability. This is
61
particularly important for uranium (U), which often persists in contaminated groundwater and
62
sediments as the soluble uranyl cation (UO22+) containing the oxidized U(VI) species. Its
63
solubility facilitates the spread of the contaminant plume far away from the source (12) and
64
results in volumes of contaminated groundwater and sediments too large to permit standard
65
‘excavation and removal’ and ‘pump and treat’ remediation approaches (13). Hence, a
66
promising bioremediation approach would be to immobilize the U(VI) contaminant and reduce it
67
to the U(IV) species, which is less soluble and can precipitate out of solution under the right
68
redox conditions and pH (12). This prevents the migration of the contaminant, reduces its
3
69
bioavailability, and minimizes the potential for exposure to humans and other living components
70
of the ecosystem.
71
One way to reduce U(VI) to U(IV) is by the action of some dissimilatory metal-reducing
72
bacteria, which can gain energy for growth by coupling the oxidation of various electron donors
73
to the reduction of the uranyl cation (14-16). This metabolic ability can be stimulated in situ with
74
field-scale additions of electron donors and results in the removal of the uranyl cation from the
75
groundwater and its accumulation in the sediments as sparingly soluble, less mobile U(IV)
76
minerals (17-21). Field-scale additions of electron donors often stimulate the growth and activity
77
of dissimilarity metal-reducing microorganisms within the family Geobacteraceae (17, 19, 20,
78
22). Studies with the model representative Geobacter sulfurreducens indicate that the reduction
79
of U by these organisms is extracellular; this prevents the permeation and reductive
80
precipitation of the radionuclide inside the cell envelope, thereby protecting the cell from its
81
toxicity (23). The extracellular reduction of U by G. sulfurreducens is primarily mediated by hair-
82
like protein filaments or pili and, to a lesser extent, by c-cytochromes of the outer membrane
83
(23). The pili of Geobacter are conductive (24) and their expression levels are linearly
84
proportional to the amount of U reductively precipitated by planktonic cells (23). Pili expression
85
in G. sulfurreducens is also required to form an electroactive biofilm matrix (25, 26), where the
86
cells are encased in a exopolysaccharide matrix containing c-cytochromes (27). This suggests
87
that Geobacter biofilms may be particularly suitable to catalyze the speciation of U while
88
providing a microenvironment that also protects cells from its toxicity.
89
Biofilms are particularly prevalent at the matrix-well screen interface and within rock fractures
90
in subsurface environments contaminated with U, thereby influencing its mobility and speciation
91
(13). However, the contribution of Geobacter biofilms to these processes and their potential use
92
as bioremediation tools has not been fully investigated. This contrasts with the availability of
93
studies about U transformations mediated by Geobacter planktonic cells (14, 15, 23, 28, 29) or
94
by biofilms formed by other metal-reducing bacteria such as Desulfovibrio (30, 31) and
4
95
Shewanella (32, 33). Hence, we investigated U transformations mediated by biofilms of G.
96
sulfurreducens. The results indicate that the biofilms have enhanced capacity to immobilize and
97
reduce U compared to planktonic cells and also tolerate exposure to higher concentrations of
98
the contaminant for prolonged periods of time. Furthermore, microscopic, genetic, and
99
spectroscopic studies of the U-reducing biofilms indicated that the radionuclide was reduced
100
extracellularly in a catalytic process influenced by the biofilm structure and the presence of
101
redox components of the biofilm matrix, most significantly the conductive pili. These findings
102
support the notion that Geobacter biofilms contribute to the immobilization and reduction of U in
103
the subsurface and highlight their potential use as permeable biobarriers for the in situ
104
bioremediation of U.
105 106
MATERIALS AND METHODS
107
Strains and culture conditions. Wild-type (WT) Geobacter sulfurreducens PCA (ATCC
108
51573), a pilin-deficient mutant (24) (herein designated pilA), and its genetically complemented
109
strain pRG5::pilA (24) (herein designated pilA+) were routinely cultured in fresh water (FW)
110
medium (34) modified as previously described (23) and supplemented with 15 mM acetate and
111
40 mM fumarate (FWAF). The medium was dispensed into tubes or serum bottles, sparged with
112
N2:CO2 (80:20), sealed with butyl rubber stoppers (Bellco) and aluminum tear-off seals
113
(Wheaton), and autoclaved 30 minutes. Unless otherwise indicated, incubations were at 30 oC.
114
Biofilm cultures. Biofilms were either grown on 6-well, cell-culture-treated polystyrene
115
plates (Corning) or on glass coverslips assembled vertically, four at a time, onto rubber
116
stoppers, as described previously (26). Briefly, the glass coverslips used for the stopper
117
assemblies were first acid-washed overnight in a bath of 50/50 (v/v) HCl/NO3- or 15% (v/v)
118
HCl/H2O and rinsed thoroughly with ddH2O before inserting them into slits cut on inverted
119
stoppers. Each coverslip assembly was placed upside down (coverslips down) in a sterile 50 ml
120
conical tube (Corning) and submerged in FW medium lacking vitamins, minerals, acetate, and
5
121
fumarate before sterilizing it by autoclaving, as previously described (26). The assembly was
122
then transferred to a clean conical tube. The biofilm chambers (either wells of plates or conical
123
tubes with coverslip assemblies) were filled with 6 or 20 ml of FWAF medium, respectively, and
124
inoculated to a final OD600 of 0.04 with an early stationary-phase FWAF culture. The chambers
125
were incubated at 30 °C for 24, 48, or 72 h, as specified, inside a vinyl glove bag (Coy Labs)
126
with a H2:CO2:N2 (7:10:83) atmosphere. At the end of the incubation period, the culture
127
supernatant fluids were decanted from the biofilm vessels and the biofilms were washed once
128
with sterile, anaerobically-prepared wash buffer (29).
129
When indicated, the biofilm biomass in the samples was estimated as total cell protein.
130
To do this, the biofilm biomass was scraped off from the surface with a sterile plastic spatula,
131
harvested by centrifugation (5 min, 12,000 x g), and the biofilm cells were then lysed in 2M
132
NaOH at 100 oC for 1 h. The lysate solution was allowed to cool before neutralizing it with an
133
equal volume of 2M HCl. After another cycle of centrifugation to remove cellular debris, the
134
supernatant, which contained the soluble protein released after lysing the cells, was analyzed
135
for protein content using a Pierce Microplate BCA Protein Assay Kit (reducing reagent
136
compatible, Thermo Scientific) with BSA standards, according to the manufacturer’s
137
specifications. Protein was measured as an OD562 on a Tecan Sunrise Plate Reader (Tecan,
138
Inc.).
139 140
U immobilization assays with resting planktonic cell suspensions and biofilms.
141
The ability of planktonic or biofilm cells to immobilize U was assayed by monitoring the removal
142
of U(VI), provided as uranyl acetate (1 mM, or as indicated), at 30 oC by resting planktonic cells
143
and biofilms, using protocols adapted from those described previously (23, 29). Planktonic cells
144
were harvested from mid-exponential cultures (OD600, 0.3-0.5) grown for 48 h under pili-inducing
145
conditions (25 oC) and suspended in 10 ml of reaction buffer (OD600, 0.1) as described before
146
(23). Resting biofilms were prepared by first growing the surface-attached communities in
6
147
FWAF medium for 24, 48, or 72 h on the coverslip-stopper assemblies described above. The
148
culture broth of the biofilm culture was then decanted gently and the coverslip assembly was
149
rinsed gently with sterile, anaerobically-prepared wash buffer (29). To initiate the assay, the
150
assemblies were submerged upside down in 50-ml conical tubes containing 20 ml of reaction
151
buffer (23, 29) supplemented with 20 mM sodium acetate and 1 mM uranyl acetate (Electron
152
Microscopy Sciences), which had been prepared from stock solutions in 30 mM bicarbonate
153
buffer. Heat-killed planktonic cells (23) and uninoculated biofilm controls were also included to
154
rule out any abiotic removal activity and/or absorption. Resting planktonic cells, resting biofilms,
155
and the controls were incubated for up to 24 h at 30 °C. Supernatant samples (500 l) were
156
periodically removed during incubation, filtered (0.22 µm Millex-GS filter, Millipore), acidified in
157
2% nitric acid (500 l), and stored at -20 °C. All procedures were performed inside an anaerobic
158
glove bag, as described above. The concentration of U(VI) in the acidified samples was
159
measured using a Platform Inductively Coupled Plasma Mass Spectrometer (ICP-MS)
160
(Micromass,Thermo Scientific) or a Kinetic Phosphorescence Analyzer (KPA) (Chemchek) and
161
provided a measure of the total U immobilized in each sample. For comparisons, the amount of
162
U immobilized by the planktonic and biofilm cells was also normalized by the cell biomass,
163
which was calculated as the amount of total cell protein released from the samples after lysis
164
with 2M NaOH (100 oC for 1 h) and neutralization with 2M HCl, as described above.
165 166
U valence and speciation by X-ray Absorption Spectroscopy (XAS) analyses. The
167
valence and speciation of the U immobilized by resting biofilms was estimated by XANES (X-ray
168
Absorption Near Edge Spectroscopy) and EXAFS (Extended X-ray Absorption Fine Structure
169
Analysis), respectively. For these analyses, biofilms grown on coverslip-stopper assemblies and
170
exposed to U for 24 h, as described above, were rinsed gently with wash buffer, scraped off the
171
coverslips, and suspended in 2 ml of reaction buffer. The biofilm biomass was then harvested
7
172
by centrifugation (12,000 x g, 10 min), loaded into custom-made plastic holders, and stored at -
173
80 °C (23). All procedures were carried out in an anaerobic chamber and samples were kept
174
frozen during XAS measurements. The XAS measurements were performed with a
175
multielement Ge detector in fluorescence mode using the PNC-CAT beamline 20-BM at the
176
Advanced Photon Source (Argonne National Laboratory) and standard beamline parameters, as
177
described elsewhere (35). XANES measurements were used to calculate the fraction of the
178
immobilized U that had been reduced to U(IV) by linear combination fitting of the spectrum with
179
U(VI) and U(IV) standards. The spectra were energy aligned using a simultaneously measured
180
uranyl nitrate standard. The U LIII-edge EXAFS spectrum was also collected for the biofilm
181
samples and modeled to determine the atomic coordination about U, as previously described
182
(23). The number of axial oxygen (Oax) atoms calculated in the EXAFS spectral analyses was
183
also used to estimate the amount of U(VI) and U(IV) in the samples, as there are two double-
184
bonded Oax atoms for each U(VI) atom in the uranyl cation (Oax=U=Oax) and none for U(IV)
185
(36).
186 187
Vitality fluorescent assays. The respiratory activity of 48-h old biofilms after exposure
188
to various concentrations (1, 2.5, and 5 mM) of uranyl acetate for 24 h was assayed using the
189
fluorescent RedoxSensor vitality kit (Invitrogen) in reference to biofilms not exposed to uranyl
190
acetate (0 mM controls), with protocols adapted from those described previously (23). The
191
biofilms were first grown for 48 h on coverslip assemblies, rinsed gently with reaction buffer, and
192
incubated in reaction buffer in the presence or absence of the uranyl acetate. The experiment
193
also included negative controls (0* mM) in which the 0 mM biofilms were first treated for 5 min
194
with electron transport chain uncouplers (a mix of sodium azide [10 mM] and carbonyl cyanide
195
3-chlorophenylhydrazone or CCCP [10 μM]) provided in RedoxSensor vitality kit (Invitrogen).
196
After 24 h of incubation, the reaction buffer was decanted from the tubes and the assemblies
8
197
were washed once with wash buffer. The biofilm biomass was scraped from the coverslips and
198
suspended in 1 ml of reaction buffer, vortexed briefly, mixed 1:1 with the Redox dye solution,
199
and incubated at room temperature for 10 min before measuring the dye’s fluorescence (490
200
nm excitation, 520 nm emission) using a SpectraMax M5 plate reader (Molecular Devices).
201
Separate aliquots of the biofilm samples were stained with SYTO 9 (Invitrogen) following
202
manufacturer’s recommendations to determine the total biofilm biomass. The respiratory activity
203
of the biofilms was estimated as the vitality index, i.e., the relative fluorescence emission from
204
the RedoxSensor dye normalized by the fluorescence emission from the Syto9 dye for 6
205
replicate samples (3 biological samples and two technical replicates for each). Statistically
206
significant (p < 0.05, one star; p < 0.005, two stars) changes in vitality indexes were determined
207
in pairwise comparisons to the 0 mM control biofilms using the t-test function of the Microsoft
208
Excel software.
209 210
Microscopy. An Scanning Electron Microscope (SEM) was used to image 48-h old WT
211
biofilms, which were grown on coverslip assemblies as described above except that round (12-
212
mm diameter), rather than square, glass coverslips were used. The biofilms were incubated at
213
30 oC in reaction buffer in the presence or absence of 1 mM uranyl acetate for 24 h and washed
214
once in wash buffer before fixing the cells at 4°C for 1-2 h in 4% glutaraldehyde. After this, the
215
biofilms were rinsed briefly in 0.1 M sodium phosphate buffer and then dehydrated in a series of
216
ethanol washes (25%, 50%, 75%, 95%, 10 minutes each) followed by three 10-min washes in
217
100% ethanol. The samples were critical point dried using a Blazers 010 critical point dryer
218
(Blazers Union Ltd.) with liquid CO2 as the transitional fluid. Once dry, the coverslips containing
219
the biofilm samples were mounted on aluminum stubs using epoxy glue and coated with ~10
220
nm of osmium using a NEOC-AT osmium coater (Meiwafosis Co., Ltd.). Samples were imaged
9
221
with a JEOL JSM-7500F SEM equipped with an Energy Dispersive Spectroscopy (EDS) 30mm2
222
detector crystal, which was used for elemental analyses.
223
Confocal Laser Scanning Microscopy (CLSM) was also used to examine the biofilms
224
and generate 3D-images for structural analyses. The WT biofilms were grown for 24, 48, and 72
225
h on 6-well plates and the pilA and pilA+ biofilms were grown for 48 h. At the end of the
226
incubation period, the culture supernatant fluid from each well was carefully decanted, and the
227
biofilms were stained for 15 min with the LIVE/DEAD® BacLight™ Bacterial Viability Kit
228
(Invitrogen) dye solution, following the manufacturer’s recommendations. After staining, the
229
biofilms were washed once in phosphate buffer saline (PBS) and imaged on a Zeiss Pascal
230
LSM microscope (Carl Zeiss Microscopy, LLC) equipped with an Achroplan 40x/0.80W
231
submersible objective. Images were collected every 1.14 µm, and side and top projections of
232
the biofilms were created using the Zeiss LSM Image Browser software (Carl Zeiss Microscopy,
233
LLC). Approximately 6-10 distinct fields of view (1,024 x 1,024 pixels, 0.22 µm/pixel) were
234
imaged for each of three independent biofilm replicates and the micrographs were analyzed with
235
the COMSTAT image analysis software using connected volume filtration to remove noise in the
236
data, as described previously (4). The biofilm structural parameters quantified with the
237
COMSTAT analyses are listed in Table S2.
238 239
SDS-PAGE and heme-stain of proteins of the biofilm EPS matrix. The
240
exopolysaccharide matrix (EPS) of 48-h biofilms grown on 6-well plates was extracted using a
241
modification of previously described protocols (27, 37). Briefly, biofilms were scraped off the
242
wells, collected in reaction buffer, and the solution was centrifuged for 10 min at 13,000 x g.
243
After decanting the supernatant fluid, the biofilm pellet was suspended in 1/5 volume of TNE (10
244
mM Tris-HCl [pH 7.5], 100 mM NaCl, 5 mM EDTA) and vortexed for 1 min. SDS was then
245
added to a final concentration of 0.1% (w/v), and the solution was mixed at room temperature
246
for 5 min. The samples were then passed 10 times through an 18G needle, and centrifuged at
10
247
15,500 x g for 20 min to collect the insoluble sheared biological fraction. The resulting pellet,
248
which predominantly contained the biofilm matrix and associated proteins, was washed 5 times
249
to remove any SDS before suspending it in 10 mM Tris-HCl buffer (pH 7.5).
250
Heme-containing proteins in the EPS biofilm matrix were also analyzed, as previously
251
described (27). The protein content in the matrix sample was determined with the bicinchoninic
252
acid (BCA) protein assay, as described above, and the equivalent of 20 µg of protein of EPS
253
sample was boiled in SDS sample buffer for 10 min before electrophoretic analysis in a 12%
254
Mini-Protean TGX gel (BioRad) at 250 V for 30 min. The Novex Sharp markers (Invitrogen)
255
were used as molecular weight standards. Heme-containing proteins were visualized on the gel
256
after N,N,N’,N’-tetramethylbenzidine staining, as described previously (23, 38). A duplicate gel
257
was run in parallel and stained for total protein using Coomassie Brilliant Blue G-250 (BioRad)
258
according to the manufacturer’s recommendations.
259 260
RESULTS AND DISCUSSION
261
Enhanced U immobilization and tolerance by biofilms. The kinetics of U
262
immobilization were investigated by measuring the amount of U, provided as uranyl acetate,
263
removed from solution by resting 48-h old biofilms and in reference to pili-expressing planktonic
264
cells (Fig. 1A). Heat-killed planktonic cell controls and uninoculated biofilm vessel controls were
265
also included to rule out any abiotic precipitation of U either due to components of the medium
266
(such as phosphate) or adsorption to the biofilm coverslip assemblies and vessels used for the
267
assays (Fig. 1A). The kinetics of U removal by the biofilms was linear throughout the 24-h long
268
assay (~10 µM/h; R² = 0.967). By contrast, the planktonic cells stopped removing U from
269
solution after 12 h and all the U immobilized by the planktonic cell biomass was solubilized
270
again after 24 h of incubation. The enhanced capacity of the biofilms to immobilize U compared
271
to planktonic cells (Fig. 1A) cannot be explained by differences in cell biomass because the
272
rates of U immobilization per h, once normalized by the total cell protein, were 6-times higher in
11
273
the biofilms (~2.8 µmol U immobilized per mg protein per h) than in planktonic cells (~0.45
274
µmol/mg/h). Furthermore, the biofilms sustained the rates of U immobilization 2 times longer (24
275
h) than their planktonic counterparts. As a result, the yields of U immobilization by the biofilms at
276
the end of the 24-h assay were 12-times greater (~66 µmol of U immobilized per mg of biofilm
277
biomass protein) than the maximum yields measured in the planktonic cells after 12 h (~5.4
278
µmol of U per mg of protein).
279
The sustained immobilization of U by 48-h old biofilms for 24 h suggests that the biofilm
280
cells remained viable and metabolically active throughout the assay. To investigate this, we
281
used the fluorogenic RedoxSensorTM dye to measure the respiratory activities of the biofilms
282
after exposure to increasing concentrations of U (1, 2.5, and 5 mM) for 24 h and in reference to
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biofilms incubated similarly but without uranium (0 mM) (Fig 1B). Interestingly, the vitality index
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of the biofilms exposed to 1 mM U was greater than the unexposed 0 mM controls.
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Furthermore, increasing the concentrations of U (2.5 and 5 mM) did not decrease the
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respiratory activities of the biofilms to or below unexposed biofilm levels. The measured vitality
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correlated well with the respiratory activity of the biofilms because pretreating the biofilms with
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sodium azide and CCCP (0* mM controls), which are chemicals that uncouple the electron
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transport chain, reduced the vitality by 80%. The dye used in these experiments yields green
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fluorescence when modified by the bacterial reductases, which are mostly located in the
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electron transport system of the cell envelope (39, 40). It is unlikely that the increases in the
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biofilm’s respiratory rate with U were due to a cellular response of the cells to U toxicity,
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because the vitality index also serves as a proxy of viability and overall cell health in G.
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sulfurreducens cells after U exposure (23). For example, exposure of pili-expressing planktonic
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cells to 1 mM U for only 6 h reduces the respiratory activity of the cells by 70%, because of the
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toxic effects of the U that permeates and mineralizes in the cell envelope (23). Yet when the
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cells are encased in the protective biofilm microenvironment they tolerated high, otherwise toxic
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levels of U, and their metabolic activities were stimulated as well.
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U reduction by the biofilms. As the reaction of interest during U bioremediation is the
300
reduction of the soluble U(VI) species to the less mobile U(IV), we used XANES to investigate if
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any of the U immobilized by the biofilms had been reduced to U(IV). For these experiments, the
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48-h old biofilms studied in Fig. 1A were collected at the end of the 24-h U challenge and the
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valence of the biofilm-associated U was analyzed by XANES in reference to U(VI) and U(IV)
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standards. Approximately 65% (± 5%) of the U immobilized by the biofilms was in the reduced
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state, U(IV) (Table S1). Consistent with the reductive precipitation of U by the biofilms, SEM-
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EDS analyses revealed needle-like, extracellular precipitates coating the microcolonies (Fig. 2A)
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that contained U (Fig. S1). The preferential localization of U minerals on the top regions of the
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biofilm is consistent with the metabolic stratification of cells within electroactive biofilms of G.
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sulfurreducens (41). As the biofilms grow, their thickness increases and a gradient of acetate is
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formed across the biofilm. This concentrates the metabolically active cells in the top regions of
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the biofilm, where acetate is more available. As a result of this metabolic stratification, U is
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preferentially reduced in the metabolically-active, top biofilm stratum (41).
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SEM micrographs also revealed the presence of extracellular filaments interspersed with
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the U precipitates (Fig. 2B). The dense filament network was more clearly visible in biofilm
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controls not exposed to U (Fig. 2C). Some filaments had diameters (< 4 nm) that closely
316
matched the diameters reported for the conductive pili of G. sulfurreducens (24), whereas
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others had larger diameters (ca. 15-20 nm) within the ranges reported for dehydrated EPS
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fibers (27). The conductive pili of G. sulfurreducens are the primary sites for extracellular
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immobilization and reduction of U by pili-expressing planktonic cells (23) and are also required
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for the formation of electroactive biofilms (25, 26). In addition to containing the conductive pili,
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the EPS matrix of G. sulfurreducens also anchors several c-cytochromes involved in metal
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reduction (27). Thus, the Geobacter matrix provides an extended matrix for U immobilization but
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also contains redox-active components which could mediate its reduction.
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U immobilization and reduction during biofilm development. As the biofilms
325
develop, their thickness increases and more biofilm matrix and matrix-associated redox
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components are available to immobilize and reduce U, respectively. Hence, we hypothesized
327
that older and thicker biofilms would immobilize and reduce more U than younger and thinner
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biofilms. For this reason, we studied U immobilization and reduction as a function of biofilm age
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(24-h, 48-h, and 72-h old biofilms) (Table S1). The biofilms were also imaged by confocal
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microscopy (Fig. 4) and the confocal micrographs were analyzed with the COMSTAT software
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(4) to estimate the thickness and other structural parameters of the biofilms (Table S2).
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We first studied U immobilization during biofilm development. Although the capacity of
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the biofilms to immobilize U increased steadily as they aged (Fig. 3A) and their thickness
334
increased (Fig. 4), the amount of U immobilized per biomass unit did not change significantly
335
during biofilm development (Fig. S3A). For example, young (24-h old) biofilms had less biofilm
336
biomass, thickness, and surface area (cells per biofilm layer) than older (48-h and 72-h old)
337
biofilms (Fig. S4, panels A-C). Yet despite being less dense, they immobilized more U per
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biofilm biomass unit than older biofilms (Fig. S3A). Interestingly, surface coverage by all of the
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biofilms was similar and neared confluence (81-92%) (Fig. S4D). The surface to volume ratio
340
also remained relatively constant during biofilm development (Fig. S4E), indicating that the area
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of the biofilm exposed to the liquid millieu relative to the biofilm volume is the same. This also
342
maintained the biofilm topography constant, as indicated by the similar roughness coefficients
343
measured in all the biofilms (Fig. S4F). Taken together, the results support a model in which
344
achieving biofilm confluence early on in biofilm development is critical to promote U
345
immobilization. As the biofilms age, they adapt their structure to maintain a topography that
346
maintains sufficient areas of the biofilms exposed to the liquid milleu and maintained similar
347
levels of U immobilization per biomass unit. This model also implies that U is preferentially
348
immobilized in the biofilm regions exposed to the liquid milley, which is further supported by the
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preferential localization of U on the outer layers of the biofilms revealed in SEM micrographs
350
(Fig. 2).
351
The fraction of the immobilized U that was reduced to U(IV) was also estimated by
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XANES (Table S1). Interestingly, the yields of U reduction increased linearly with biofilm age
353
(Fig. 3A), even when normalized by the biofilm biomass (Fig. S3A). None of the biofilm
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structural parameters determined in the COMSTAT analyses fit the linear trend of U reduction
355
(Fig. S4). Parameters important for U immobilization (surface coverage, surface to volume
356
ration, and roughness coefficient) were, for example, unaffected by biofilm age. The biofilm
357
parameters that changed were those related to biofilm biomass (biofilm biomass, thickness, and
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surface area) but they all stabilized in the 48-h and 72-h old biofilms rather that increasing
359
linearly. By contrast, the total protein content of the biofilms, which accounts for protein
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contributed by cells and by proteinaceous components of the biofilm matrix, increased linearly
361
as the biofilms aged (Fig. S5). Biofilm formation is a developmental process, consisting of
362
specific stages such as attachment, microcolony formation, and biofilm maturation (42). As they
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transition from one stage of biofilm development to the next, the biofilms increase their
364
thickness and biomass but also undergo extensive gene reprogramming so that specific biofilm
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components and activities are expressed (9, 43). Pili expression in G. sulfurreducens is, for
366
example, required to transition from young, monolayered biofilms to thicker biofilms composed
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of several layers of cells (26) and this, in turn, is required to maintain the electroactivity of the
368
biofilms (25). Hence, as the Geobacter biofilms age, they express more redox-active
369
components in the biofilm matrix and this, in turn, may increase their capacity to reduce the
370
immobilized U to U(IV).
371 372
Role of the redox components of the biofilm matrix in U immobilization and
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reduction. As the conducive pili of G. sulfurreducens are the primary site for U immobilization
374
and reduction in planktonic cells (23) and also are required for biofilm formation (25, 26), we
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compared the capacity of 48-h biofilms of the WT, a pilin-deficient mutant (pilA), and its
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genetically complemented strain pilA+ to immobilize and reduce U (Fig. 3B). The pilA biofilms
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had a diminished capacity to immobilize and reduce U (p = 0.07) compared to WT biofilms of
378
the same age. The defect was due to the pilus deficiency because complementing the mutation
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in trans (pilA+) restored the phenotypes and promoted U immobilization and reduction to levels
380
comparable to the WT biofilms.
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The defect in pili production in the pilA biofilms also prevented the biofilms from growing
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in thickness (Fig. 4) and lead to ~ 50% decreases in other biomass-related structural
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parameters such as biomass volume and surface area (cells per biofilm layer) compared the
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WT biofilms (Table S2 and Fig. S4A-C). These defects are consistent with the reported role of
385
the pili at promoting cell-cell aggregation during biofilm formation and providing the structural
386
support required to build multilayered biofilms (26). However, despite the biomass and pili
387
defects, the pilA biofilms immobilized more U per biomass unit than the WT biofilms.
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Furthermore, genetic complementation of the pilA mutation in the pilA+ strain resulted in denser
389
biofilms (Fig. 4), consistent with the increased ability of the pilA+ hyperpiliated strains to
390
aggregate (26). Yet despite the increased biofilm biomass, the pilA+ biofilms immobilized less U
391
per biofilm biomass unit than any other biofilm (Fig. S3B). The lack of a clear correlation
392
between biofilm biomass and U immobilization in the pilA, WT, and pilA+ biofilms may reflect
393
differences in diffusion rates of electron donors and acceptors, as more diffusional constraints
394
are expected in thicker (pilA+>WT) than in thinner (pilA) biofilms. Consistent with this, the pilA+
395
biofilms had lower surface to volume ratios than any other biofilms (Fig. S4E and Table S2),
396
indicating that less surface area of the biofilm volume is exposed to the liquid medium and
397
available for electron donors and acceptors diffusion. Furthermore, the roughness coefficient,
398
which provides a measure of the spatial heterogeneity of the biofilms as variations in biofilm
399
thickness, was also lowest in the pilA+ biofilms (Fig, S4F). This indicates that the very dense
400
pilA+ biofilms have a relative uniform topography (very low roughness coefficient, ~0.06),
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401
whereas the thin pilA biofilms formed the most heterogenous biofilms of all (highest roughness
402
coefficient, ~0.32, almost twice the WT coefficients) (Table S2). Hence, the more heterogenous
403
structure of the pilA and WT biofilms also maximizes the biofilm surface area available for
404
diffusion of electron donors and acceptors.
405
Although the pili-deficient pilA biofilms immobilized more U per biomass unit than any
406
other strain, they reduced a lower fraction (~38%) of the immobilized U to U(IV) than the WT
407
(~63%) and pilA+ (~76%) biofilms per biomas unit (Fig. S3). The capacity of the biofilms to
408
reduce U (pilA
