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Characterization of uraninite nanoparticles produced byShewanella oneidensis MR-1

William D. Burgos a,*, Jeffrey T. McDonough a, John M. Senko a, Gengxin Zhang a,Alice C. Dohnalkova b, Shelly D. Kelly c, Yuri Gorby d, Kenneth M. Kemner c

a Department of Civil and Environmental Engineering, The Pennsylvania State University, 212 Sackett Building, University Park,

PA 16802, USAb Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory, Richland, WA, USA

c Biosciences Division, Argonne National Laboratory, Argonne, IL, USAd J. Craig Venter Institute, La Jolla, CA, USA

Received 6 December 2007; accepted in revised form 25 July 2008; available online 6 August 2008

Abstract

The reduction of uranium(VI) by Shewanella oneidensis MR-1 was studied to examine the effects of bioreduction kinetics andbackground electrolyte on the physical properties and reactivity to re-oxidation of the biogenic uraninite, UO2(s). Bioreductionexperiments were conducted with uranyl acetate as the electron acceptor and sodium lactate as the electron donor under restingcell conditions in a 30 mM NaHCO3 buffer, and in a PIPES-buffered artificial groundwater (PBAGW). MR-1 was cultured inbatch mode in a defined minimal medium with a specified air-to-medium volume ratio such that electron acceptor (O2) limitingconditions were reached just when cells were harvested for subsequent experiments. The rate of U(VI) bioreduction was manip-ulated by varying the cell density and the incubation temperature (1.0 � 108 cell ml�1 at 20 �C or 2.0 � 108 cell ml�1 at 37 �C) togenerate U(IV) solids at ‘‘fast” and ‘‘slow” rates in the two different buffers. The presence of Ca in PBAGW buffer altered U(VI)speciation and solubility, and significantly decreased U(VI) bioreduction kinetics. High resolution transmission electron micros-copy was used to measure uraninite particle size distributions produced under the four different conditions. The most commonprimary particle size was 2.9–3.0 nm regardless of U(VI) bioreduction rate or background electrolyte. Extended X-ray absorp-tion fine-structure spectroscopy was also used to estimate uraninite particle size and was consistent with TEM results. The reac-tivity of the biogenic uraninite products with dissolved oxygen was tested, and neither U(VI) bioreduction rate nor backgroundelectrolyte had any statistical effect on oxidation rates. With MR-1, uraninite particle size was not controlled by the bioreductionrate of U(VI) or the background electrolyte. These results for MR-1, where U(VI) bioreduction rate had no discernible effect onuraninite particle size or oxidation rate, contrast with our recent research with Shewanella putrefaciens CN32, where U(VI) bio-reduction rate strongly influenced both uraninite particle size and oxidation rate. These two studies with Shewanella species canbe viewed as consistent if one assumes that particle size controls oxidation rates, so the similar uraninite particle sizes produced byMR-1 regardless of U(VI) bioreduction rate would result in similar oxidation rates. Factors that might explain why U(VI) bio-reduction rate was an important control on uraninite particle size for CN32 but not for MR-1 are discussed.� 2008 Elsevier Ltd. All rights reserved.

1. INTRODUCTION

The United States’ history of utilizing uranium resourcesfor weapons and energy production has left an environmen-

tal hazard that poses a present and future threat to theintegrity of soil and groundwater (Riley and Zachara,1992). Uranium in contaminated aquifers is most prevalentin a soluble VI oxidation state (Langmuir, 1978). Microbi-ologically mediated reduction of U(VI) to a relativelyinsoluble U(IV) (UO2(s)—uraninite) is a strategy for immo-bilizing U (Langmuir, 1978; Taylor, 1979), and may be

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doi:10.1016/j.gca.2008.07.016

* Corresponding author. Fax: +1 814 863 7304.E-mail address: wdb3@psu.edu (W.D. Burgos).

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stimulated in contaminated subsurface environments by theaddition of a physiological electron donor (Lovley et al.,1991). However, this strategy is only effective if the U(IV)remains reduced, since U(IV) may be oxidized back toU(VI) resulting in continued mobility of U (Finneran etal., 2002; Sani et al., 2005; Senko et al., 2005; Ginder-Vogelet al., 2006). In order for enzymatic uranium reduction tobe implemented, the potential oxidation of immobilized Uprecipitates should be considered.

A factor that could contribute to the stability of U afterit has been bioreduced is the nature of the U(IV) precipi-tates. Biogenic U(IV) phases may range in size from mono-meric U(IV) (Suzuki et al., 2002; Boyanov et al., 2007) toindividual UO2 (uraninite) nanoparticles ranging in sizefrom 0.9 to 5 nm (Suzuki et al., 2002; Sani et al., 2005; Mar-shall et al., 2006; Senko et al., 2007) to large aggregates ofup to 100 nm (Abdelouas et al., 1999). Similarly sized ura-ninite nanoparticles have been produced through the abi-otic reduction of U(VI) with sulfide (Beyenal et al., 2004)and green rust (O’Loughlin et al., 2003). The cellular loca-tion, particle size and aggregation state of biogenic U(IV)precipitates undoubtedly influence their susceptibility tooxidative remobilization (Beller, 2005; Marshall et al.,2006; Senko et al., 2007) and the potential for transportof nanoparticulate U(IV) in groundwater (Suzuki et al.,2002). Indeed, work by Fredrickson et al. (2002) suggestedthat extracellularly deposited U(IV) produced by Shewanel-

la putrefaciens CN32 could be oxidized by Mn(III/IV) oxi-des, while periplasmic deposition of U(IV) could ‘‘protect”it from reaction with Mn(III/IV) oxides. Furthermore, lar-ger, highly aggregated U(IV) nanoparticles (produced viarelatively slow rates of U(VI) bioreduction by CN32) weremore resistant to oxidation than relatively small, and lessaggregated U(IV) nanoparticles (produced via relatively ra-pid rates of U(VI) reduction), suggesting that the rate ofU(VI) reduction influences the nature of U(IV) particlesand therefore their susceptibility to re-oxidation (Senko etal., 2007).

The rate of U(VI) reduction is by no means the only fac-tor controlling the nature of biogenic U(IV) precipitates.The geochemical conditions under which U(VI) reductiontakes place, and the physiological state of the microorgan-isms mediating U(VI) reduction (among other factors) willundoubtedly exert control on the nature of biogenic U(IV)phases. For example, the presence of dissolved calcium mayinhibit U(VI) bioreduction (Brooks et al., 2003), and cal-cium incorporation into U(IV) minerals (Burns, 1999)could influence their susceptibility to re-oxidation. Shewa-

nella species are a relatively well-characterized group ofU(VI)-reducing bacteria that contain a complex and appar-ently redundant system of electron transport for the reduc-tion of U(VI) and other metals (e.g., Fe(III), Mn(III/IV))(Wade and DiChristina, 2000; Beliaev et al., 2001; Shi etal., 2006; Bretschger et al., 2007). This redundancy may ex-plain the retention of some metal-reducing activity and thedifferences in cellular location of U(IV) phases (i.e., peri-plasmic or extracellular) upon deactivation of genes whoseproducts are known to reduce U(VI) (Marshall et al., 2006).Furthermore, the conditions under which Shewanella spe-cies are cultured appear to strongly influence outer mem-

brane composition and excretion of extracellularcomponents (DiChristina et al., 2002; Fang et al., 2006;Ruebush et al., 2006; Biju et al., 2007; Neal et al., 2007),some of which are redox-active proteins (Marshall et al.,2006) and components of electrically conductive bacterialnanowires (Gorby et al., 2006). Therefore, the conditionsunder which a microorganism is cultured could influencethe production and location of U(VI)-reducing enzymes,which, in turn, could influence the cellular location, particlesize and aggregation state of U(IV) precipitates.

In light of these observations, we assessed the nature ofU(IV) precipitates produced by S. oneidensis MR-1 grownunder carefully controlled conditions (Hill, 2007) designedto ‘‘poise” that organism for the production of bacterialnanowires that are believed to play a significant role inthe reduction of U(VI) and other metals (Gorby et al.,2006). This is in contrast to previous experiments in whichU(IV) precipitates were produced via U(VI) reduction by S.

putrefaciens CN32 that was grown in a complex medium(Senko et al., 2007). We manipulated rates of U(VI) reduc-tion by S. oneidensis MR-1 in bicarbonate buffer and anartificial groundwater by varying the cell density and theincubation temperature. Biogenic U(IV) precipitates werecharacterized by X-ray absorption spectroscopy (XAS)and electron microscopy. The reactivity of these U(IV) pre-cipitates was characterized by their susceptibility to oxida-tion by dissolved oxygen.

2. EXPERIMENTAL SECTION

2.1. Background electrolytes

All experiments were conducted in one of two back-ground electrolytes: 30 mM NaHCO3 (pH 6.8, preparedunder an 80:20% N2:CO2 atm; referred to hereafter asNaHCO3 buffer) or a 10 mM Piperazine-N,N0-bis-(2-eth-anesulfonic acid) (PIPES)-buffered artificial groundwater(pH 6.3, prepared under an 85:15% N2:CO2 atm; referredto hereafter as PBAGW buffer). Both buffers also included5 mM sodium lactate and 1.2–1.6 mM uranyl(VI) acetate.The components of PBAGW (aside from PIPES) were se-lected to simulate the groundwater from Area 2 at theOak Ridge National Laboratory Field Research Center(Brooks, 2001). The concentrations of all components ofPBAGW buffer are presented in electronic annex EA-1.Critical component concentrations of PBAGW with respectto uranyl speciation included 5 mM NaHCO3, 4.2 mMCaCl2, and 0.01 mM KH2PO4. The PBAGW buffer alsocontained macronutrients for bacterial growth whereasthe NaHCO3 buffer did not.

2.2. Cell cultivation

Shewanella oneidensis MR-1 (referred to hereafter asMR-1) was cultured in a chemically defined medium similarto PBAGW except with the addition of vitamins, trace met-als (Tanner, 1997), arginine, glutamine, serine (Myers andNealson, 1988), and ferric(III) nitrilotriacetic acid, and withthe exclusion of uranyl acetate (electronic annex EA-1). ThePBAGW-defined growth medium was prepared under oxic

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conditions, pH adjusted to 6.8–7.1 with NaOH, and dis-pensed into serum bottles (165 ml average total volume).Fifty-four milliliter of medium were added to each bottleand autoclaved, then 1 ml of filter-sterilized ferric NTA(5 mM) was added to establish a 2:1 air:liquid ratio (vol:-vol) in all bottles. A frozen cell suspension of MR-1 wasthawed, transferred to a culture tube containing TrypticSoy Broth-without dextrose (TSB-D) and incubated aerobi-cally for 24 h at 30 �C and 50 rpm. This culture was used toinoculate the PBAGW defined growth medium, which wasthen incubated in the sealed bottles for 15.5 h at 30 �C and50 rpm. Under these conditions, oxygen in the headspacelimited growth (Hill, 2007). Cell growth reached late-logphase at ca. 16 h. TEM images of MR-1 cells at differentincubation times revealed that these culture conditions in-duced the production of extracellular appendages that grewin size and abundance between 16 and 24 h (electronic an-nex Fig. EA-1-1). An incubation time of 15.5 h was selectedto harvest cells before this extracellular material wasproduced.

2.3. U(VI) bioreduction experiments

Bioreduction experiments were conducted in 1-L Pyrexbottles equipped with anoxic sampling caps. An anoxiccap consisted of a Balsh tube (bottom cut off) insertedthrough a cored thick black stopper fitted into the bottletop and secured with a plastic screw cap (center hole re-moved), and the protruding Balsh tube was sealed with athick butyl stopper and aluminum crimp top. Reactors werefilled with ca. 500 ml of anoxic buffer (either NaHCO3 orPBAGW) containing 5 mM sodium lactate, uranyl acetatestock solution was added to achieve an initial U(VI) con-centration of 1.2–1.6 mM, and reactors were purged witha N2:CO2 gas mix corresponding to the buffer. Experimentswere initiated by the addition of washed cell suspensions.Cell density was measured by absorbance at 600 nm andcorrelated to cell number by acridine orange direct counts.The A600 value at the time of cell harvest was 0.182 ± 0.021based on over 500 trials and was continually validated.Cells were centrifuged (15 min at 3500g and 20 �C) underanoxic conditions and resuspended and washed three timesin anoxic buffer (either NaHCO3 or PBAGW). Cell densityand incubation temperature were varied to manipulate therates of U(VI) bioreduction. With either buffer, ‘‘fast” bio-reduction conditions were achieved using an MR-1 densityof approximately 2.0 � 108 cell ml�1 maintained at 37 �C,and ‘‘slow” bioreduction conditions were achieved usingan MR-1 density of 1.0 � 108 cell ml�1 maintained at20 �C. All reactors were kept in darkness with no mixing ex-cept for sample collection. After cells were added, sampleswere periodically removed with sterile needle and syringe(in anoxic chamber) and soluble and total U(VI) concentra-tions were quantified as described below.

2.4. U(IV) oxidation experiments

All biogenic uraninite oxidation experiments were con-ducted in 30 mM NaHCO3-buffered solution (pH 6.8, un-der 80:20% N2:CO2 atm) regardless of the buffer used in

the previous bioreduction incubation, and were conductedin 120 ml serum bottles sealed with thick butyl stoppersand aluminum crimp tops. After a bioreduction incubationtime of 30–50 days, whereupon soluble U(VI) was removedfrom solution by MR-1 to less than 19–33 lM U(VI) underthe four conditions, cell-uranium precipitates were pasteur-ized (70 �C for 30 min) to deactivate biological activity. Nogrowth was observed when pasteurized cell-uranium precip-itates were used to inoculate TSB-D. Cell-uranium precipi-tates were added to NaHCO3 buffer to achieve a total Uconcentration of 150–200 lM (initially predominantlyU(IV)). Oxygen was provided to incubations by flushingthe headspace of anoxic NaHCO3 buffer-containing serumbottles with filter-sterilized air. To maintain the pH of thebicarbonate buffered medium, 20% of the air headspacewas removed and replaced with an equivalent volume ofCO2. No change in medium pH resulted from thesemanipulations.

2.5. U(VI) measurements

Soluble U(VI) was measured after centrifugation for10 min at 14,100g and 20 �C. Total U(VI) (including solubleand solid-associated U(VI)) and total U (including U(VI)and U(IV)) were measured as described by Elias et al.(2003). Briefly, samples of well-mixed suspensions were re-moved from incubations by syringe in an anoxic glovebag(95:5% N2:H2 atm; Coy Laboratory Products Inc., GrassLake, MI) and placed in 1 M anoxic NaHCO3 (pH 8.4).After extraction for 5 min, solids were removed by centrifu-gation and U(VI) was measured in the supernatant. Tomeasure total U, suspension samples were placed in oxic10% HNO3 under air for at least 12 h to oxidize allU(IV). U(VI) in all operational procedures was measuredby kinetic phosphorescence analysis on a KPA-11 (Chem-Chek Instruments, Richland, WA) (Brina and Miller,1992).

2.6. Electron microscopy

For SEM, samples were prepared in a glove box follow-ing a previously published procedure (Zhang et al., 2007).Briefly, cell-mineral suspensions were fixed in anoxic 2.5%glutaraldehyde, placed on a glass cover slip, and mineralparticles were allowed to settle onto the cover slip for15 min. The particle-coated cover slips were graduallydehydrated in an ethanol series followed by critical pointdrying (CPD). All sample preparation, except CPD, wasperformed in an anoxic glovebag to minimize the exposureof samples to O2. Cover slips were mounted onto a SEMstub and Au coated for observation using a Zeiss Supra35 FEG-VP SEM at an accelerating voltage of 10–15 kV.A short working distance (6–10 mm) and low beam current(30–40 mA) were used to achieve the best image resolution.A longer working distance (8.0 mm) and higher beam cur-rent (50–70 mA) were used for qualitative energy dispersivespectroscopy (EDS) analysis. For whole mount TEM, cellswere fixed in 4% glutaraldehyde and placed on a 400 meshcarbon-sputtered formvar-coated copper grid. After 5 min,excess suspension was removed, the cells were negatively

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stained with 2% uranyl acetate, air dried and examined witha JEOL 1200 EX II TEM at 80 kV. For thin-section TEM,all sample preparation was performed in a Coy anoxicglovebag. Cell-uranium precipitates were fixed in anoxic2.5% glutaraldehyde followed by 3 washes in anoxic water,gradual dehydration in an ethanol series and infiltration inLR White resin. Polymerized samples were sectioned to70 nm on a microtome (Leica Ultracut UCT), and sectionswere mounted on 200 mesh copper grids coated with form-var support film sputtered with carbon. Sections wereexamined using JEOL 2010 high resolution transmissionelectron microscope (HR TEM) equipped with LaB6 fila-ment operating at 200 kV with a resolution of 1.9 A

0. Ele-

mental analysis was performed using an Oxford EDSsystem equipped with a SiLi detector coupled to theTEM, and analyzed with ISIS software. Images were digi-tally collected using a Gatan CCD camera and analyzedusing Gatan Digital Micrograph. d-spacings of selectedarea diffraction (SAED) ring patterns were evaluated byDesktop Microscopist software.

2.7. X-ray absorption spectroscopy

XAS measurements were made at the Materials Re-search Collaborative Access Team (MR-CAT) sector 10-ID beamline (Segre et al., 2000) of the Advanced PhotonSource at Argonne National Laboratory. The XAS spectrawere collected in transmission mode using quick-scanningof the monochromator. X-ray absorption near edge struc-ture (XANES) spectra from the cell-uranium precipitateswere used to determine the average valence state withinthe samples. The X-ray absorption edge energy was cali-brated by collecting the reference spectrum from hydrogenuranyl phosphate (U(VI) Std) during the collection of eachspectrum. All data sets were accurately aligned in energyusing the derivative of the edge of the U(VI) standard. Alinear combination fitting (LCF) of the U(VI) Std and abiogenic nanoparticulate UO2 standard (U(IV) Std)(O’Loughlin et al., 2003) was used to determine the approx-imate U(VI) to U(IV) ratio in the sample XANES spectra.This approach assumes that the standard spectra representthe uranyl species in the samples. This assumption can leadto a systematic uncertainty for these samples because we donot know the U(VI) species. As a result we used our solidU(VI) phosphate standard and an aqueous uranyl(VI) car-bonate standard, as likely U(VI) components of the sam-ples. The valence state of the samples was verified byusing the number of axial oxygen atoms of the uranyl(Oax) as determined by analysis of the EXAFS data. Thismethod is less model-dependent because it does not assumeprior knowledge of the uranyl species.

At the end of the bioreduction incubation period, celluranium-precipitates were stored at 4 �C before shipmentto ANL. Samples for XAS were referred to their bufferand bioreduction condition as follows: NaHCO3 ‘‘fast”,NaHCO3 ‘‘slow”, PBAGW ‘‘fast”, and PBAGW ‘‘slow”.Samples (approximately 40 mg cell-uranium precipitates)were mounted as moist pastes on filter paper or as centri-fuged pastes mounted in a plastic holder, covered withKapton film and sealed with Kapton tape. All sample prep-

aration was performed in a Coy anoxic glovebag and allsamples were stored in the chamber prior to analysis. Sam-ples mounted in the holders were exposed to the atmo-sphere for less than 1 min before being mounted for XASmeasurements in a free-flowing N2 environment to limitpossible sample oxidation. These XAS sample holders havebeen shown to maintain anoxic integrity, when exposed toan oxic environment, for at least 8 h.

Uranium XANES spectra were collected in 30-s inter-vals consecutively for 5 min. No radiation-induced changesto the XANES spectra were observed at the 30-s intervals ofdata collection. Extended X-ray absorption fine-structure(EXAFS) spectra were collected in 5-min intervals, aver-aged and then the background was removed using the pro-grams Athena (Ravel and Newville, 2005) and IFEFFIT(Newville, 2001). The background removal parameter Rbkgwas set to 1.0 A (Newville et al., 1993). The effect of double-electron excitations like those described by Hennig (2007)were removed from these spectra as described elsewhere(Ravel and Kelly, in press). The EXAFS spectra were mod-eled using theoretical models from atomic clusters of urani-nite (Wyckoff, 1960) as input for FEFF 7.02 (Zabinskiet al., 1995). The EXAFS parameters were optimized tothe measured spectra using the program FEFFIT (Sternet al., 1995). The EXAFS parameters are the amplitudereduction factor ðS2

0Þ, an energy shift to align the theoreticaland measured spectra (DE0), the number of atoms within ashell or coordination number (CN), distance between theuranium atom and the neighboring atoms (R), and themean square displacement in that distance (r2). Details ofthe EXAFS model are provided in electronic annex EA-2.

3. RESULTS

3.1. U(VI) bioreduction kinetics

The rate of U(VI) bioreduction by MR-1 was observedto be first-order with respect to the soluble U(VI) concen-tration (Fig. 1) according to:

Rred ¼ d½UðVIÞ�=dt ¼ �kred � ½UðVIÞðsolÞ� ð1Þ

where kred is the first-order bioreduction rate constant(d�1). The rate of U(VI) bioreduction was certainly affectedby other factors such as lactate concentration, cell/U(VI)ratio, cell/lactate ratio, temperature and (non-)mixing con-ditions but these factors are purposefully excluded from Eq.(1) for a simplified comparison. As designed, differencesamong the rates of U(VI) bioreduction were dependenton cell concentration and temperature (Table 1). For exam-ple, kred increased by a factor of 24 in NaHCO3 buffer whenthe cell concentration was doubled and the incubation tem-perature increased by 17 �C. Similarly, but to a lesser ex-tent, kred increased by a factor of 7.6 in PBAGW bufferbetween the ‘‘fast” and ‘‘slow” bioreduction conditions.Liu et al. (2002) reported a first-order U(VI) bioreductionrate constant of 28.4 d�1 for MR-1 in experiments con-ducted with 0.50 mM uranyl chloride, 5 mM Na-lactate,and 2.0 � 108 cell ml�1 at 30 �C in 30 mM NaHCO3 buffer.While this is reasonably close to our reported rate constantof 11 d�1 for the ‘‘fast” condition in NaHCO3 buffer, the

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difference could have been caused by differences in cell cul-turing techniques (e.g., MR-1 grown on TSB vs. PBAGWdefined growth medium with finite oxygen in headspace)and/or temperature effects (e.g., optimal growth tempera-ture for MR-1 is 30 �C).

The U(VI) bioreduction rate constants obtained inPBAGW buffer were 3.5 to 11 times less than rate constantsobtained in NaHCO3 buffer under otherwise identical con-ditions. The slower rate of U(VI) bioreduction in PBAGWbuffer was likely caused by important differences in U(VI)speciation and solubility. The presence of Ca has a signifi-cant effect on the rate and extent of U(VI) bioreduction be-cause of the decreased reduction potential of U(VI)–Cacomplexes as compared to U(VI)–CO3 complexes (Brookset al., 2003; Stewart et al., 2007). The bioreduction rate ofsolid-phase U(VI) has been shown to be slower than soluble

U(VI) because of limited bioavailability caused by dissolu-tion kinetics and/or bioenergetic costs required to utilize asolid-phase electron acceptor (Fredrickson et al., 2000).Using Visual MINTEQ (Gustafsson, 2008) to model ura-nium speciation in the NaHCO3 buffer with 1 mM U(VI)at pH 6.8, we found the predominant U(VI) species wereUO2ðCO3Þ4�3 (86% of total) and UO2ðCO3Þ2�2 (12%) and>99% of the U(VI) would remain in solution. Abiotic con-trols for the NaHCO3 buffer showed no difference betweentotal and soluble U(VI) concentrations and no decline inthese concentrations over the incubation period. UsingVMINTEQ to model uranium speciation in PBAGW bufferwith 1 mM U(VI) at pH 6.3, the predominant soluble U(VI)species was Ca2UO2(CO3)3 (40% of total), 45% of theU(VI) was predicted to remain in solution, and the predom-inant U(VI) solid phase was rutherfordine (UO2CO3, 54%

0

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Fig. 1. U(VI) bioreduction kinetics by Shewanella oneidensis MR-1 in (a and c) NaHCO3 buffer and (b and d) PBAGW buffer. With eitherbuffer, ‘‘fast” bioreduction conditions were achieved using a cell concentration of 2.0 � 108 cell ml�1 maintained at 37 �C, and ‘‘slow”

bioreduction conditions were achieved using a cell concentration of 1.0 � 108 cell ml�1 maintained at 20 �C. Measured triplicate data(symbols) and first-order rate models (lines).

Table 1First-order rate constants for U(VI) bioreduction (kred, Eq. (1)) by Shewanella oneidensis MR-1, and subsequent first-order rate constants foruraninite oxidation (kox, Eq. (2)) by dissolved O2

Sample name kred (d�1) r2 P-value kox (d�1) r2 P-value

NaHCO3 ‘‘fast” 11 0.97 <0.010 16 0.88 <10�4

NaHCO3 ‘‘slow” 0.45 0.87 <10�4 27 0.79 <0.011PBAGW ‘‘fast” 0.99 0.98 <10�4 17 0.98 <10�4

PBAGW ‘‘slow” 0.13 0.86 <10�3 16 0.78 <10�3

Sample names refer to the buffer and bioreduction condition during U(VI) reduction.

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of total). Uninoculated controls for the PBAGW buffershowed consistent differences between total and solubleU(VI) concentrations over the incubation period, withinsoluble U(VI) accounting for 21–30% of the U(VI). Thus,the decreased solubility of U(VI) and the predominance ofthe Ca2UO2(CO3)3 species in PBAGW buffer are believedto have decreased the rate of bioreduction by MR-1. De-tails of the uranium speciation modeling are provided inelectronic annex EA-3.

We have operationally defined the concentration of so-lid-associated U(VI) as the difference between the totaland soluble U(VI) concentrations, and the percentage of so-lid-associated U(VI) as this quantity divided by the initialtotal U(VI) concentration. At the end of the bioreductionexperiments in NaHCO3 buffer, the percentage of solid-associated U(VI) was 17.0 ± 0.65 and 8.59 ± 1.78% in the‘‘fast” and ‘‘slow” conditions, respectively. This residualfraction of U(VI) was likely incorporated into the biogenicuraninite particles as reported by others (Conradson et al.,2005). At the end of the bioreduction experiments inPBAGW buffer, the percentage of solid-associated U(VI)was 15.3 ± 0.77 and 21.7 ± 2.23% in the ‘‘fast” and ‘‘slow”

conditions, respectively. The residual U(VI) in PBAGWbuffer was likely distributed between the initial unreducedU(VI) solid-phases (primarily rutherfordine predicted byVMINTEQ) and biogenic uraninite particles.

3.2. Electron microscopy of cell-uranium precipitates

SEM images of biogenic uraninite precipitates producedby MR-1 in NaHCO3 buffer suggested that solids tended toaccumulate extracellularly (Fig. 2) consistent with recentwork to establish the cellular location of U(VI) reductasesand biogenic U(IV) (Marshall et al., 2006). The greateramount of solids observed under ‘‘slow” conditions waslikely caused by the higher U/cell ratio used in these exper-

iments (i.e., more uraninite generated per cell). Uraniniteparticles tended to associate with and accumulate on extra-cellular polymeric substances (EPS) (Fig. 2a–d), perhapselectrically conductive biological nanowires (Gorby et al.,2006). SEM images of biogenic uraninite particles producedby MR-1 in PBAGW buffer suggested that solids tended toprecipitate directly on the cell surface resulting in thicklycoated cells (Fig. 2e and f). If one assumes that U(VI) solidsformed during the incubation period due to rutherfordinesupersaturation, then the cells could have been in directcontact with a solid-phase electron acceptor.

TEM images of uraninite precipitates produced by MR-1 in either buffer were consistent with their correspondingSEM images (Figs. 3 and 4). In NaHCO3 buffer, the major-ity of uraninite particles were located outside of the cell,slight amounts of uraninite were located within the peri-plasm under ‘‘fast” conditions and even less amounts under‘‘slow” conditions (Fig. 3). The connectivity of uraniniteparticles along EPS was apparent in many images but rep-resents only a 2-dimensional view of what are likely com-plex, 3-dimensional features, similar to those seen inprevious studies (Marshall et al., 2006). In PBAGW buffer,uraninite precipitated almost exclusively on the outer mem-brane under ‘‘fast” conditions, while uraninite precipitatedboth in the periplasm and on the outer membrane under‘‘slow” conditions (Fig. 4). The exceptionally thick coatingson the outer membranes and the large deposits located out-side of the cells (Fig. 4b and d) resembled the lipopolysac-charide layer of Shewanella spp. observed by Korenevskyet al. (2002).

The size of the uraninite particles were measured by highresolution TEM. In samples with thick uraninite coatingsor large extracellular deposits, identifying what constitutesa single, discrete particle was challenging. We have definedthe primary particle as the smallest cohesive unit in theseimages, versus larger aggregates of these units. We have

Fig. 2. Scanning electron micrographs of unstained Shewanella oneidensis MR-1 cells and uraninite precipitates after near-complete removalof soluble U(VI) in (a–c) NaHCO3 buffer under ‘‘slow” bioreduction conditions, (d) NaHCO3 buffer under ‘‘fast” bioreduction conditions, (e)PBAGW buffer under ‘‘fast” bioreduction conditions, and (f) PBAGW buffer under ‘‘slow” bioreduction conditions. (a–c) Provide a series ofimages of increasing magnification that detail extracellular polymeric substances.

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chosen this unit because, from a thin-section perspective,the size of this primary particle is least likely to be affectedby the plane-angle sectioned through the sample material.For example, with a larger polymerized aggregate, dimen-sions of uraninite particles observed in one view may notbe observed in another sectioned plane. Particle size distri-butions, based on 56 measurements from different areaswithin one section for each experimental condition, showedvery similar trends (Table 2; histograms in electronic annexFig. EA-4-1). The most common uraninite primary particlesize was 2.9–3.0 nm regardless of bioreduction rate or back-ground electrolyte.

While we have defined particle size based on the primaryparticles, we observed aggregated particles where d-spacinglines extended across two or more particles, especially in thethick coatings produced in PBAGW buffer under ‘‘slow”

conditions. For the PBAGW ‘‘slow” sample, the size of60 aggregates that showed extended d-spacing lines weremeasured. A histogram of these data showed a sharp peakat ca. 6 nm, essentially a doubled value of the primary par-ticle size measured under all conditions (TEM images andhistogram provided in electronic annex EA-4). The continu-ity of d-lines between uraninite nanoparticles presentedhere is similar to those between ferrihydrite nanoparticlesduring transformation of ferrihydrite to goethite (Banfieldet al., 2000; Burleson and Penn, 2006), and such oriented

aggregation-dependent crystal growth may be controlledby biopolymers (Nesterova et al., 2003).

3.3. X-ray absorption spectroscopy of cell-uranium

precipitates

XANES and EXAFS were used to quantify the uraniumoxidation state in the bioreduced cell-uranium precipitates.The results from both methods (linear combination fittingof XANES spectra, and number of uranyl axial oxygenatoms from the EXAFS spectra) were consistent (Table 3).The uncertainties based on the EXAFS method are largerthan those based on the XANES method and are the valueswe present here, because, as discussed in the methods sec-tion, the EXAFS method is less dependent on the assumedspeciation in the samples. The XANES for cell-uraniumprecipitates produced in PBAGW and NaHCO3 buffers clo-sely matched that of a U(IV) standard of biogenic UO2

nanoparticles, although there is a small shift in the spectra,indicating a component of U(VI) (Fig. 5). This U(VI) com-ponent was also identified during analysis of the EXAFSdata (Fig. 6). EXAFS analysis of the cell-uranium precipi-tates produced in NaHCO3 buffer under ‘‘fast” and ‘‘slow”

conditions contained 80 ± 9% and 90 ± 7% U(IV), respec-tively, with the remainder as U(VI). EXAFS analysis ofthe cell-uranium precipitates produced in PBAGW buffer

Fig. 3. Transmission electron micrographs of unstained Shewanella oneidensis MR-1 cells and uraninite precipitates after near-completeremoval of soluble U(VI) in NaHCO3 buffer under (a and c) ‘‘fast” bioreduction conditions, and (b and d) ‘‘slow” bioreduction conditions.Notice heavy extracellular precipitation of uraninite preferentially associated with extracellular polymeric substances (b and d).

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under ‘‘fast” and ‘‘slow” conditions contained 86 ± 6% and89 ± 8% U(IV), respectively. XAS-based uranium oxida-tion states were in good agreement with our operationaldefinition for solid-phase U(VI) based on extractions andKPA analysis as listed in Table 3.

Cell-uranium precipitates were characterized by EXAFSto determine uranium speciation (Fig. 6). For these cell-ura-

nium precipitates, EXAFS spectra were modeled with acombination of uranyl as described by 2 Oax atoms andan equatorial oxygen shell (O1), and uraninite as describedby an oxygen shell (O1), a U shell (U), and a more distantoxygen shell (O2). The percentage of U(VI) in the sampleswas determined by a weighting factor multiplied by 2 Oaxatoms of the uranyl (Kelly et al., 2008). It is possible that

Fig. 4. Transmission electron micrographs of unstained Shewanella oneidensis MR-1 cells and uraninite precipitates after near-completeremoval of soluble U(VI) in PBAGW buffer under (a and c) ‘‘fast” bioreduction conditions, and (b and d) ‘‘slow” bioreduction conditions.Cells in ‘‘fast” conditions showed slight uraninite deposition (a and c), in contrast to the ‘‘slow” conditions. Cells in ‘‘slow” (b and d)experiment feature periplasmic and outer membrane specific deposition of nanocrystalline uraninite.

Table 2Average uraninite particle size based on EXAFS coordination number for U neighbors, and based on HR TEM measurements

Sample name Number of U neighborsa Particle sizeb (nm) Particle sizec (nm)

Lower bound Best-fit Upper bound Mean ± sd

NaHCO3 ‘‘fast” 9.5 ± 0.9 2.0 2.7 3.6 3.03 ± 0.45NaHCO3 ‘‘slow” 10.1 ± 1.2 2.2 3.4 4.5 2.85 ± 0.31PBAGW ‘‘fast” 7.5 ± 1.0 1.0 1.2 2.1 3.01 ± 0.26PBAGW ‘‘slow” 8.1 ± 0.7 1.2 1.5 2.2 2.96 ± 0.21

Sample names refer to the buffer and bioreduction condition during U(VI) reduction.a Corrected for the percentage of U(VI) in the sample.b Size based on EXAFS fits. EXAFS loses it sensitivity for large particle sizes, therefore, the uncertainties are smaller for the lower bound

than for the higher bound.c Size based on 56 discrete particles measured for each sample using HR TEM.

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the U(VI) in these samples was part of the uraninite struc-ture (Conradson et al., 2005), a hydrated species, an ab-sorbed species that is weakly associated with a ligand(Kelly et al., 2002), or a precipitated rutherfordine mineral.However, no additional statistically significant signal wasneeded to describe the measured EXAFS spectra of thesesamples. Thus, the uranyl species could not be defined.For example, attempts to fit the U EXAFS data, via theinclusion of a U–C scattering path, consistent with thetwo bidentate coordinated carbonates associated with U

in rutherfordine, resulted in U–C coordination numbersof 0.2–0.4 with an uncertainty of 0.5. If all of the U(VI)in the PBAGW samples (11–14% as identified in Table 3)existed as rutherfordine, U–C coordination numbers of0.22–0.28 would be expected. Thus, although the EXAFSresults are consistent with 100% of the U(VI) within thePBAGW samples existing as rutherfordine, because of thelarge uncertainties of the EXAFS coordination number val-ues, the EXAFS results are also consistent with 0% of theU(VI) within these same samples existing as rutherfordine.Similar scenarios resulted with all attempts to fit the datawith a U(VI) component associated with uraninite. There-fore, although the analysis of the XANES and EXAFSspectra do identify a U(VI) component within all of thesamples, they do not provide conclusive evidence as tothe identity of the uranyl species within them.

The U–U coordination number of the uraninite particlesdetermined by EXAFS was used to estimate average urani-nite particle sizes (Table 2). This coordination number wasadjusted for the fraction of U(VI) in the sample. For bio-genic uraninite produced in NaHCO3 buffer under ‘‘fast”and ‘‘slow” bioreduction conditions, the average uraniniteparticle sizes were estimated to be approximately2.7 ± 0.8 and 3.4 ± 1.2 nm, respectively. For biogenic ura-ninite produced in PBAGW buffer under ‘‘fast” and ‘‘slow”

bioreduction conditions, the average uraninite particle sizes

Table 3Average valence state within cell-uranium precipitates as deter-mined by wet chemical methods (extractions and KPA analysis),linear combination fitting (LCF) of the XANES spectra, and fromthe number of axial oxygen atoms determined from the EXAFSspectra

Sample name % U(IV) in precipitates

KPA analysis LCF XANES EXAFS

NaHCO3 ‘‘fast” 83.0 ± 0.7 95 ± 1 80 ± 9NaHCO3 ‘‘slow” 91.4 ± 1.8 75 ± 1 90 ± 7PBAGW ‘‘fast” 84.7 ± 0.8 83 ± 3 86 ± 6PBAGW ‘‘slow” 78.3 ± 2.2 85 ± 3 89 ± 8

Sample names refer to the buffer and bioreduction condition duringU(VI) reduction.

Fig. 5. U LIII-edge normalized XANES spectra (symbols), linear combination fit (red line), and the U(VI) and U(IV) components of the fit for(a) NaHCO3 ‘‘fast”, (b) PBAGW ‘‘fast”, (c) NaHCO3 ‘‘slow”, and (d) PBAGW ‘‘slow”.

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were estimated to be approximately 1.2 ± 0.8 and 1.5 ±0.8 nm, respectively. There is a slight trend of larger particlesizes with slower reduction rates, but this change is wellwithin the uncertainty of the measurements. Uraninite par-ticle size is smaller in PBAGW buffer than NaHCO3 buffer,although the uncertainties overlap. The average particle sizedetermined by EXAFS from all four samples was2.2 ± 1.0 nm, quite consistent with the average particle sizeof 3.0 ± 0.1 nm obtained from all TEM measurements.

3.4. Oxidation of bioreduced uranium precipitates

Cell-uranium precipitates were pasteurized and incu-bated with dissolved oxygen to measure their susceptibility

to re-oxidation (Fig. 7). The rate of uraninite oxidation wasquantified as the rate of soluble U(VI) production accord-ing to:

Rox ¼ d½UO2�=dt ¼ �kox � ð½UTOT� � ½UðVIÞðsolÞ�Þ ð2Þ

where kox is the first-order oxidation rate constant (d�1)and [UTOT] is the total uranium concentration (bothU(IV) and U(VI)) in the reactor. The rate constant for airoxidation of uraninite particles produced in NaHCO3 buf-fer was greater for precipitates formed under ‘‘slow” versus‘‘fast” reduction kinetics (Table 1). However, because therewas substantial overlap of the data points and only a minordifference between the fitted curves for Eq. (2), a statisticaldifference between the air oxidation rates could not be

Fig. 6. U LIII-edge EXAFS spectra from the cell-uranium precipitates (a), magnitude of Fourier transform (b), real part of Fourier transform(c), and a representation of the uraninite structure (d). The measured spectra (symbols) are shown with the models (solid lines). These setshave been displaced vertically for clarity. The samples shown from top to bottom in (a–c) are PBAGW ‘‘fast”, PBAGW ‘‘slow”, NaHCO3

‘‘fast”, and NaHCO3 ‘‘slow”. The components of the model for the PBAGW ‘‘slow” spectrum are shown at the bottom of (b and c).

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established. In fact, there were no statistical differences inthe air oxidation rates for any of the four solids tested, sug-gesting that neither bioreduction rate nor background elec-trolyte had any measurable effect on subsequent re-oxidation. This result is in contrast to our recent reporton U(VI) reduction by S. putrefaciens CN32 and subse-quent re-oxidation of biogenic uraninite (Senko et al.,2007).

4. DISCUSSION

4.1. Size and cellular location of uraninite nanoparticles

The size of uraninite particles were measured by bothhigh resolution TEM and EXAFS (Table 2 and electronicannex Fig. EA-4-1). With TEM, the measured particle sizeaveraged 3.0 ± 0.1 nm for all conditions which is consistentwith the average best-fit EXAFS particle size of 2.2 ±1.0 nm. EXAFS particle sizes are commonly smaller thanthose obtained by microscopy methods, in part, becausethe EXAFS-based measurement represents an average ofthe sizes of particles in the sample that includes monomericU(IV) and particles too small to be visualized by TEM.Thus, a larger discrepancy between TEM and EXAFS par-ticle size results in PBAGW buffer can be explained by themore complicated solution chemistry that could possibly

stabilize more monomeric U(IV). Importantly, consideringaccuracy of and differences in techniques, all particle sizemeasurements were in agreement.

Uraninite nanoparticles were located in the periplasm,on the outer membrane, and extracellularly, consistent withanother recent study with MR-1 (Marshall et al., 2006).TEM images have been used to interpret the predominanceof any particular U(VI) reduction mechanism. On onehand, the observation of densely packed precipitates inthe periplasm suggests the involvement of periplasmic cyto-chromes in U(VI) reduction (Fredrickson et al.,2000, 2002;Senko et al., 2007). On the other hand, observation of pre-cipitates on the outer membrane suggests that cytochromeslocalized on the outside of the outer membrane are involvedin U(VI) reduction (Myers and Myers, 1992; DiChristina etal., 2002). Finally, observation of precipitates along EPS orglycocalyx suggests the involvement of extracellular cyto-chromes in U(VI) reduction (Gorby et al., 2006; Marshallet al., 2006). These observations underscore the apparentredundancy of electron transport in Shewanella species.

4.2. Geomicrobiological implications

Presently, the bioremediation of several important metaland radionuclide contaminants (e.g., Cr(VI), Tc(VII) andU(VI)) is based on the reduction of soluble species topoorly soluble minerals. The long-term success of this ap-proach hinges on the reactivity (ideally, extremely limited)of these reduced minerals with oxidants that may eventuallyenter post-biostimulated environments. The chemical reac-tivity of these reduced minerals will be controlled by a num-ber of factors including biophysical characteristics of theprecipitates (e.g., particle size, aggregation state, cellularlocation), geochemical characteristics of the infiltrating oxi-dizing fluids (e.g., specific oxidant, oxidant concentration,solution chemistry), and geophysical characteristics of theaquifer material (e.g., heterogeneity, location of precipitateswith respect to fluid flow paths, temperature). We have spe-cifically attempted to discern how bioreduction rate andbackground electrolyte affect biophysical characteristics ofbiogenic uraninite.

In our previous study with Shewanella putrefaciens

CN32 (Senko et al., 2007) we reported a U(VI) bioreduc-tion rate formulation that was zero-order with respect to[U(VI)]. For comparison of U(VI) bioreduction rates fromthat study to those of the present study, we calculated pseu-do first-order rate constants for U(VI) reduction accordingto Eq. (1). The data from Senko et al. (2007) fit a pseudofirst-order rate formulation nearly as well as a zero-orderrate formulation, and pseudo first-order rate constants ofU(VI) reduction by Shewanella putrefaciens CN32 (Table4) were quite similar to those of Shewanella oneidensis

MR-1 (Table 1). For example, the first-order U(VI) biore-duction rate constant for experiments conducted at 23 �Cwith 9.1 � 107 cells CN32 ml�1 in 30 mM NaHCO3 was0.24 d�1 (r2 = 0.92), while the first-order U(VI) bioreduc-tion rate constant for experiments conducted at 20 �C with1.0 � 108 cells MR-1 ml�1 in 30 mM NaHCO3 was 0.45 d�1

(r2 = 0.87). Therefore, we feel these two studies are kineti-cally similar and provide insight into the relationships be-

Fig. 7. Oxidation kinetics of biogenic uraninite particles bydissolved oxygen. Uraninite precipitates produced by Shewanella

oneidensis MR-1 in (a) NaHCO3 buffer or (b) PBAGW buffer. Themeasured data (symbols) are shown with the first-order rate models(lines).

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tween bioreduction rate, uraninite particle size and re-oxi-dation rate. With MR-1, uraninite particle size was not con-trolled by the bioreduction rate of U(VI) or the backgroundelectrolyte. Assuming that particle size controls re-oxida-tion, because all uraninite particles were of similar size,the subsequent uraninite oxidation rates were essentiallyconstant. With CN32, uraninite particle size was controlledby the bioreduction rate of U(VI). Because a wider range ofuraninite particle sizes were produced (smallest particlesproduced at fastest rates), the subsequent uraninite oxida-tion rates varied significantly (smallest particles re-oxidizedfastest). These two studies with Shewanella species can beviewed as consistent if one assumes that particle size con-trols oxidation rates, so the similar uraninite particle sizesproduced by MR-1 regardless of U(VI) bioreduction ratewould result in similar oxidation rates.

By considering our results with both MR-1 and CN32 itis apparent that bioreduction rate is not the only factor thatcontrols primary particle size of biogenic uraninite. A num-ber of other inter-related factors could control particle size,including: cell cultivation methods, metabolic state of cells,molecular-scale mechanisms of U(VI) reduction, U(IV)nucleation site, cellular location of uraninite precipitates,and secondary effects of oriented aggregation. Cell cultiva-tion methods may have been a critical difference betweenour studies with MR-1 and CN32. For instance, cultivatingcells on a minimal medium induces different expression pat-terns than cultivation on a rich, complex medium (Tao etal., 1999). MR-1 was cultured in sealed bottles of a definedminimal medium such that cells were harvested at the onsetof electron acceptor (O2) limiting conditions (Hill, 2007).Electron acceptor-limiting conditions have been shown topromote the formation of outer membrane cell surface do-mains on MR-1 that contain heme proteins (Biju et al.,2007). As with Fe(III) reductase activity in MR-1 (Beliaevet al., 2005; Ruebush et al., 2006), U(VI) reductase activityis also likely dependent on growth conditions. CN32 wasgrown through an extended period of electron acceptor-lim-iting conditions such that its metabolic state would havebeen different than the metabolic state of MR-1 used in thisstudy.

The molecular-scale mechanism of U(VI) reduction andsubsequent U(IV) precipitation could also affect uraniniteparticle size. It is currently unknown if physical featuresof a U(VI) reductase would affect the size of U(IV) mineralproducts because it is also unknown exactly where U(IV)

precipitation occurs. However, if U(VI) was reduced by aU(VI) reductase in the periplasm, the growth and aggrega-tion of uraninite could be affected by the constrained size ofthis region. On the other hand, if U(VI) was reduced by aU(VI) reductase on the outside of the outer membraneand U(IV) diffused away from the cell surface, particlegrowth may be more readily facilitated by oriented aggrega-tion of U(IV) particles. Secondary effects of oriented aggre-gation, beyond factors that control the production and sizeof primary particles, could also be important regardless ofthe physiological location of the uraninite particles. Forexample, organic molecules or extracellular exudates couldbe associated with the uraninite primary particles and affectoriented aggregation (Nesterova et al., 2003).

Other recent studies have also examined characteristicsand reactivity of biogenic uraninites produced by Shewanel-

la oneidensis MR-1 (Schofield et al., 2008), Desulfovibrio

vulgaris and Geobacter sulfurreducens (Bargar et al.,2008), and Anaeromyxobacter dehalogenans 2CP-C (Mar-shall et al., 2008). In experiments with Shewanella oneidensis

MR-1, Schofield et al. (2008) found that the EXAFS struc-tural coherence length was greater for incubations con-ducted at pH 8.0 where U(VI) was bioreduced moreslowly, as compared to incubations conducted at pH 6.3where U(VI) was bioreduced more rapidly. In addition,Bargar et al. (2008) found that inclusion of Mn2+ led tothe production of smaller uraninite particles, but that thesmaller, Mn-containing uraninite particles were more resis-tant to re-oxidation than the larger, Mn-free uraninite par-ticles. In experiments with Anaeromyxobacter dehalogenans

2CP-C, Marshall et al. (2008) found that U(VI) bioreduc-tion rates were strongly dependent on the electron donor.U(VI) bioreduction was more rapid with H2 as comparedto acetate. While the electron donor, and hence the rateof U(VI) bioreduction, did not affect UO2 nanoparticle sizeor the association of UO2 nanoparticles with extracellularpolymeric substances, donor-specific differences in UO2

nanoparticle aggregation were observed. At a relatively‘‘fast” U(VI) bioreduction rate with H2, smaller aggregateswere observed while at a relatively ‘‘slow” U(VI) bioreduc-tion rate with acetate, larger, more uniform aggregates wereobserved.

As these new findings on biogenic uraninite nanopar-ticles emerge, the complexity of biomineralization in thesesystems becomes more apparent. Clearly there are a num-ber of cellular, biochemical and geochemical variables

Table 4Comparison of U(VI) of zero-order versus first-order bioreduction rate constants for Shewanella putrefaciens CN32 from Senko et al. (2007)

Exp.condition

Zero-order rate(umol U(VI)l�1 d�1)

Zero-order rate constant(umol U(VI) l�1 d�1 (108 cell/ml)�1)

r2 N—number of datapoints in regression

First-order rateconstant (d�1)

r2 N—number of datapoints in regression

‘‘fast” 602 274 0.98 6 1.1 0.95 7‘‘medium” 182 200 0.97 8 0.24 0.92 8‘‘slow” 55 240 0.96 10 0.032 0.94 9

The zero-order rate formulation was zero-order with respect to U(VI), zero-order with respect to lactate (electron donor), and first-order withrespect to cell concentration, yielding a zero-order cell-dependent rate constant with units of (lmol U(VI) l�1 d�1 (108 cell/ml)�1). The first-order rate formulation was first-order with respect to U(VI), zero-order with respect to lactate, and zero-order with respect to cell concen-tration, yielding a first-order rate constant with units of (d�1).

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that can exert control on the size and reactivity of urani-nite nanoparticles. Regardless of our speculative discus-sion of factors that control the size of uraninitenanoparticles, oxidation experiments designed to measureuraninite reactivity were consistent with physical featuresobserved in the SEM and TEM images and spectroscopicfeatures resolved by EXAFS. This was also true with ourprevious study with CN32, where oxidation experimentsdisplayed clear differences consistent with differences inTEM images and EXAFS spectra. However, generalizedconclusions about the physical and physiological charac-teristics of biogenic precipitates related to cell metabolismrate are still difficult to make and are an area of activeresearch.

ACKNOWLEDGMENTS

This work was supported by the Environmental RemediationScience Program (ERSP), Office of Biological and EnvironmentalResearch (BER), Environmental Remediation Science Division(ERSD), US Department of Energy (DOE) Grant No. DE-FG02-04ER63914 to The Pennsylvania State University, and bythe National Science Foundation under Grant No. CHE-0431328and the U.S. Department of Energy, Biological and EnvironmentalResearch (BER). Use of the MR-CAT sector at the Advanced Pho-ton Source (APS) and of the APS was supported by the US DOE,Office of Science, Office of Basic Energy Sciences and Office of Bio-logical and Environmental Research, under contract W-31-109-ENG-38 and the MRCAT member institutions. We extend ourgratitude to Bruce Ravel and Maxim Boyanov (ANL) for theirassistance with the EXAFS data collection and to BR for his con-tribution to the interpretation and correction of the EXAFS spec-tra. Part of this work was performed at the EnvironmentalMolecular Sciences Laboratory (EMSL), a national scientific userfacility sponsored by the DOE’s OBER, located at the PacificNorthwest National Laboratory (PNNL) in Richland, WA. PNNLis operated for DOE by Battelle Memorial Institute under ContractDE-AC06-76RL0 1830.

APPENDIX A. SUPPLEMENTARY DATA

Supplementary data associated with this article can befound, in the online version, at doi:10.1016/j.gca.2008.07.016.

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Associate editor: Peggy A. O’Day

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