Uranium Redistribution Due to Water Table Fluctuations in Sandy

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Uranium redistribution due to water table fluctuations in sandy wetland mesocosms Emily R. Gilson, Shan Huang, Paul G. Koster van Groos, Kirk G. Scheckel, Odeta Qafoku, Aaron D. Peacock, Daniel I Kaplan, and Peter R Jaffe Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.5b02957 • Publication Date (Web): 25 Sep 2015 Downloaded from http://pubs.acs.org on September 25, 2015

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Environmental Science & Technology

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Uranium redistribution due to water table

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fluctuations in sandy wetland mesocosms

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Emily R. Gilson1, Shan Huang1, Paul G. Koster van Groos1†, Kirk G. Scheckel2, Odeta Qafoku3,

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Aaron D. Peacock4, Daniel I. Kaplan5, and Peter R. Jaffé1* 1

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2

6 7

3

US EPA, Cincinnati, Ohio 45268, United States

Pacific Northwest National Laboratory, Richland, Washington 99352, United States 4

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Princeton University, Princeton, New Jersey 08540, United States

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Pace Analytical Energy Services, Pittsburgh, Pennsylvania 15238, United States

Savannah River National Laboratory, Aiken, South Carolina 29808, United States

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In order to better understand the fate and stability of immobilized uranium (U) in wetland

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sediments, and how intermittent dry periods affect U stability, we dosed saturated sandy wetland

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mesocosms planted with Scirpus acutus with low levels of uranyl acetate for 4 months before

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imposing a short drying and rewetting period. Concentrations of U in mesocosm effluent

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increased after drying and rewetting, but the cumulative amount of U released following the dry

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period constituted less than 1% of the total U immobilized in the soil during the 4 months prior.

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This low level of remobilization suggests, and XANES analyses confirm, that microbial

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reduction was not the primary means of U immobilization, as the U immobilized in mesocosms

ACS Paragon Plus Environment

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was primarily U(VI) rather than U(IV). Drying followed by rewetting caused a redistribution of

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U downward in the soil profile and to root surfaces. While the U on roots before drying was

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primarily associated with minerals, the U that relocated to the roots during drying and rewetting

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was bound diffusely. Results show that short periods of drought conditions in a sandy wetland,

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which expose reduced sediments to air, may impact U distribution without causing large releases

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of soil-bound U to surface waters.

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Introduction

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Waste from uranium (U) mining and milling for energy and weapons production can

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contribute to U contamination of soils and groundwater. While dissolved in groundwater, U can

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travel from its source to contaminate surface waters used for drinking and animal habitat. For

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example, migration of U contamination has led to excessive U levels in wells, mines, and rivers

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near abandoned U mines in the southwestern United States1, 2 and in wetlands near a Department

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of Energy nuclear facility at the Savannah River Site in Aiken, SC.3 Consumption of U-

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contaminated waters can have toxic effects on the kidneys, bones, and livers of humans and

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animals.4-6

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A number of strategies have been investigated to contain and mitigate U contamination by

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limiting its mobility in groundwater. These techniques generally aim to reduce concentrations of

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U in groundwater by chemically or biologically depositing U in sediments. One of the most-

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researched of these strategies is bioreduction which relies on metal-reducing bacteria to

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immobilize U in sediments by reducing oxidized and relatively soluble U(VI) to less soluble

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U(IV).7 U immobilization through bioreduction has been demonstrated in lab-based and in situ


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systems.8-10 U can also be immobilized in situ through complexation, sorption, or coprecipitation

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of U(VI) with organic matter,11, 12 iron oxides,13 or phosphate.14

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Certain natural systems, including wetlands, have been shown to immobilize U.9,

11, 13

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Wetlands are strategically located for limiting U exposure to surface ecosystems because they

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often occur at interfaces of groundwater and surface water. Wetland biogeochemistry is thought

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to be favorable to bioreduction of U.13 Wetland sediments generally have low levels of oxygen

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which favors the anaerobic bacteria capable of U bioreduction, such as sulfate and iron (Fe)

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reducers, and which stabilizes U that has been reduced. Wetlands further promote activity of Fe-

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reducing bacteria because the organic matter originating from wetland plants acts as a carbon

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source, which supplies needed electrons, and plant roots are repositories of Fe(III) plaque, which

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bacteria like Geobacter spp. require as their typical electron acceptor.13 When U(VI) is present,

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it can complement Fe(III) as the electron acceptor in bacterial processes, but U levels are

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typically low enough that Fe(III) is also required to maintain a robust community of Fe-reducing

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bacteria. The plant-supplied organic matter in wetlands can also directly immobilize U through

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sorption.11

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The dynamic nature of wetland systems impacts the fate of the contaminants they contain.

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Wetland soils are typically saturated and anoxic, but events such as seasonal water table

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fluctuations or larger climate events like droughts may lead to more oxidizing conditions as soils

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dry and are exposed to air. Changes in biogeochemical conditions, when such fluctuations arise,

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are expected to affect the fate of U in contaminated wetlands. Reduced U(IV) is susceptible to

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reoxidation in a number of conditions including limitation of electron donors or introduction of

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oxygen, ferric hydroxides, manganese (III/IV) oxides, nitrate reduction intermediates, and

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carbonates.9, 15, 16 Should bioreduced U(IV) solids in a wetland be reoxidized to U(VI) by an


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influx of oxygen to the soil, they would likely re-dissolve and experience enhanced mobility.

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Conversely, U(VI) bound to soil components like organic matter is already oxidized and may be

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more resilient against remobilization in the event of water table fluctuations and exposure to

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oxygen.11

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The objective of this research was to determine how sediment drying and rewetting affects the

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fate of U immobilized in wetland sediments.

To study U immobilization, greenhouse

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mesocosms were built to represent an idealized sandy wetland and were dosed with uranyl

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acetate for several months. Effluent U concentrations from mesocosms were measured and

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mesocosms were dissected after being dried and rewetted. Chemical extractions, speciation

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measurements, and imaging analyses of U on sediments and roots were used to gain new insights

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into U immobilization in sandy wetlands and the effects of drying and rewetting on the stability

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of immobilized U.

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Materials And Methods

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Establishment and Maintenance of Sandy Wetland Mesocosms

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To study U immobilization in sandy wetlands, five 0.8-L mesocosms were built in a

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greenhouse. (See Supporting Information Figure S1 for sketch of mesocosm.) Four of the

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mesocosms were planted with Scirpus acutus, hardstem bulrush plants commonly found in

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wetlands, which were acquired from a local nursery (Pinelands Nursery and Supply, Columbus,

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NJ). The solid media used in the mesocosms was a 1:7.5 mixture (by weight) of sediment from

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wetlands at the Savannah River Site in Aiken, SC and acid-washed ASTM standard C778 20-30

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sand (US Silica, Ottawa, IL).

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mesocosms were consistent biogeochemically with U-contaminated wetlands and contained

The use of Savannah River Site sediment ensured that our


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bacteria capable of U reduction.3, 13 Sand was included in the mix to promote even flow of water

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through the whole mesocosm and to ease root sampling at the end of the treatment. Although the

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sediment-sand mixture deviates from real-world conditions, it is reasonably similar to sediments

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at the Savannah River Site, some of which contain very high levels of sand.3, 17 Nonetheless,

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care should be taken when applying conclusions based on this model sandy wetland to natural

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wetland systems that have high water retention, more organic carbon, and more complex

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mineralogy than this system.

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The bulrush plants were planted in a manner that permitted making biogeochemical

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measurements in sediment-sand mixtures influenced by roots and not influenced by roots.

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Rhizomes and roots of live bulrush plants were rinsed and then planted in 70-µm nylon mesh

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bags (10-cm deep by 3.5-cm radius, ~0.4 L) containing the sediment-sand mixture. The mesh

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bags were then placed in 0.8-L mesocosm buckets and backfilled with additional sediment-sand

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mixture until they were approximately 75% full and the sediment levels inside and outside the

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bags were the same. The top of the mesh bags rose approximately 3 cm above the sediment

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surface.

In the planted mesocosms, the sediments inside the bags represent rhizosphere

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sediments while the sediments outside the bags represent bulk sediments. An identical setup, but

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without plants, was used as an unplanted control mesocosm.

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deoxygenated nutrient solution at a rate of approximately 210 mL day-1. This influent solution

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was a modified Hoagland nutrient solution that contained: 1.00 mM MgCl2•6H2O, 2.00 mM

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KCl, 50.00 µM H3BO3, 0.75 µM ZnSO4•7H2O, 0.10 µM Na2MoO4•2H2O, 0.40 µM CuCl2, 0.36

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µM CaCl2•H2O, and 0.89 µM MnCl2.18

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mesocosms, 300 mM sodium acetate and 300 mM urea were added to the influent by a syringe

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pump at a rate of approximately 1.5 mL day-1 to ensure anaerobic conditions and to promote the

Mesocosms were fed a

Before the nutrient-spiked water reached the


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activity of Fe-reducing bacteria native to the Savannah River Site sediment. Water flowed up

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from basal inlets through the mesocosm sediment and exited through drains situated above the

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sediment surface after a hydraulic retention time of approximately 1.3 days. This flow scheme

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ensured that the sediment remained entirely saturated even under summer conditions when high

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light intensity and high temperatures led to increased evapotranspiration.

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temperature was not controlled and therefore varied with the season.

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temperature and time of year, the outflow rate could drop to half the inflow rate and varied

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between mesocosms. Throughout the study, plants were subjected to artificial lightning during

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the night.

Greenhouse

Depending on the

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After three to four months of growth in the greenhouse, when the plants appeared healthy with

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green shoots of approximately 1 m in length, U addition was initiated. Dissolved Fe(II) in

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mesocosm pore water varied with depth and location, but had values as high as 5.6 mg L-1,

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indicating that reducing conditions had been established by the time U addition began.

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Immediately prior to U addition, the pH values of mesocosm pore water were between 6.0 and

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6.7. To facilitate greater cumulative U immobilization, the mesocosms were operated with

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steady U addition for a long period of approximately four months. Uranyl acetate was pumped

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into the mesocosms through a separate basal inlet such that the final concentration of U in the

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two combined inflows was 9.5 µM U. Chemical equilibrium modeling using Visual MINTEQ

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ver. 3.0 software indicates that the predominant U species in the combined influents was

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(UO2)3(OH)5+ with UO2OH+ and (UO2)4(OH)7+ generally making up the second and third most

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prevalent species (see Supporting Information Table S1 for example results from Visual

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MINTEQ calculations).

These calculations represent the combined influent chemistry at


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experimental temperature and pH values but do not include possible complexation with organic

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matter.

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To determine U immobilization in the mesocosms, the differences between influent and

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effluent U concentrations were calculated. Effluent was collected from each mesocosm at

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approximately weekly intervals throughout the experiment. The tubing that drained the effluent

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from the individual mesocosms fed into a single larger tube from which samples of all the

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mesocosms’ combined effluents were also collected. Effluent samples were acidified in 2%

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HNO3 and stored at 4°C until analysis.

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Drying and Rewetting Treatments

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The drying treatment began after the mesocosms had operated under steady conditions with U

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addition for 137 days. Before the drying treatment was initiated, one of the planted mesocosms

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was dissected. (In later sections, this mesocosm is referred to as “never dried, planted (1)”.) The

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four remaining mesocosms were disconnected from the U inflow. Three of the mesocosms, one

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of which was the unplanted control, were disconnected from the water source and drained by

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pumping water out through their basal inlets. After four hours, pumping could not remove any

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additional water. Samples of this drained pore water were collected, acidified in 2% HNO3, and

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stored at 4°C until U concentrations could be analyzed. The unsaturated mesocosms remained in

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the greenhouse with no fluid inputs for 9 days. This phase of the experiment occurred during the

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month of April, thus the greenhouse temperature generally experienced daily fluctuations

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between 26°C and 30°C.

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turnover, chemical and biological kinetics, and evapotranspiration are all temperature-dependent,

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therefore U dynamics would be expected to be affected by the season.

Sorption, mineral solubility, biogeochemical reactions, carbon


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After the 9-day dry period, the water inflow was turned on in the unplanted and one of the two

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planted dry mesocosms at the same pre-drying rate of approximately 210 mL day-1. The second

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planted dry mesocosm was dissected immediately without the addition of any water. (In later

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sections, this mesocosm is referred to as “after drying, planted”.) After 2 days of slowly re-

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introducing nutrient solution, the two mesocosms were entirely re-saturated and nutrient solution

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began flowing out of the effluent drains above the sediment surface. Effluent samples were

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collected for the next 11 days to measure the U remobilized after the drying and rewetting.

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These effluent samples were acidified in 2% HNO3 and stored at 4°C until U concentrations

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could be analyzed. The two rewetted mesocosms were dissected after the 13-day rewetting

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phase. (In later sections, these mesocosms are referred to as “after rewetting, planted” and “after

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rewetting, unplanted”.)

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maintained under steady, saturated conditions with water inflow throughout the drying and

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rewetting treatments was also dissected. (In later sections, this mesocosm is referred to as “never

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dried, planted (2)”.)

The following day, the planted control mesocosm that had been

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Mesocosms were dissected before and after the drying and rewetting treatments to examine the

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treatments’ impacts on mesocosm biogeochemistry. During dissection, each mesocosm was

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placed in an anaerobic glove bag and water was pumped out of the mesocosm through one of the

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basal inlets. After approximately 4 hours, pumping could not remove any more water and the

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pump was disconnected. The nylon mesh bags holding the roots and sediments were removed

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from the mesocosms. The sediments inside the bag were separated into deep (4-7 cm below

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sediment surface) and shallow (1-4 cm below sediment surface) sections (the top 1 cm of

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sediment was discarded). The roots were similarly partitioned before they were chopped up

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using gardening shears. The bulk sediments from three horizons outside the nylon mesh bag (1-4


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cm, 4-7 cm, and 7-11 cm below the sediment surface) were separated, as well. Sediment and

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roots from each horizon were homogenized by hand before they were divided into subsamples

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that were preserved for analysis of U concentration, speciation, and distribution; Fe

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concentration; and bacterial abundance.

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Analytical Procedures

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U concentrations in acidified effluent samples were measured on an Element 2TM Inductively-

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Coupled Plasma Mass Spectrometer (ICP-MS, Thermo Scientific, Germany). Yttrium was used

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as an internal standard. Uranyl acetate standards prepared by serial dilution were used to convert

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counts per second to U concentration. U concentrations in two acidified influent samples were

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also measured by ICP-MS to confirm the calculated U concentration in the stock solution.

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Total U on sediments and roots from dissected mesocosms was extracted with bicarbonate and

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then measured by ICP-MS. Approximately 1.0 g of sediments or approximately 0.2 g of roots

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from each horizon was deposited in 5 mL of 0.2 M NaHCO3. This extraction procedure was

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adapted from previous work19 and was intended to provide a measure of the readily extractable

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sediment U concentrations, and the total plant U concentrations. Extracts were diluted and

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acidified in 2% HNO3 before U concentrations were measured on the ICP-MS.

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Total bioavailable Fe and Fe(II) on sediments and roots from dissected mesocosms were

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extracted with acid and then quantified using the ferrozine method. Approximately 1.0 g of

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sediments and approximately 0.2 g of roots from each horizon were deposited in 5 mL of 0.5 N

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HCl. After 72 hours, Fe(II) and total Fe were measured using the ferrozine method.20 Briefly,

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total Fe was measured by adding 20 µL of 6.25 M hydroxylamine hydrochloride to 1 mL of HCl-

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extracted sample. After 24 hours, all of the Fe(III) was reduced to Fe(II) which could be

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measured using the ferrozine method. 30 µL of the reduced solution was added to 1.5 mL of


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ferrozine and the absorbance was measured after 30 minutes at a wavelength of 562 nm in a

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Spectronic Genesys 2 Spectrophotometer (Thermo Electron Corporation, USA). The Fe(II)

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concentration was measured directly by adding 30 µL of HCl-extracted sample to 1.5 mL

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ferrozine and measuring the absorbance after 30 minutes at a wavelength of 562 nm in a

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Spectronic Genesys 2 Spectrophotometer (Thermo Electron Corporation, USA). Absorbances

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were converted to molar concentrations of Fe(II) using a standard curve.

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U on roots and sediments was examined to determine its oxidation state by U (LIII-edge: 17166

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eV) x-ray absorption spectroscopy (XAS) at beamline 10-ID (Materials Research Collaborative

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Access Team (MRCAT)) at the Advanced Photon Source at Argonne National Laboratory,

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Argonne, IL.21 The electron storage ring operated at 7 GeV. A liquid nitrogen cooled double

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crystal Si(111) monochromator was used to select incident photon energies and a platinum-

210

coated mirror was used for harmonic rejection. U reference materials included UO2,

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UO2(CH3COO)2•2H2O, U(UO2)(PO4)2, (UO2)8O2(OH)12•12(H2O), UO2(CO3), UO2(NO3)2, and

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U(VI) sorbed to maghemite. Freeze-dried sediment samples from all five mesocosms and freeze-

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dried root samples from the four planted mesocosms were mounted and sealed in plexiglass

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holders using Kapton tape. At least three scans were collected for each sample at room

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temperature. XAS spectra were collected in fluorescence and transmission modes using

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ionization chambers for the incident, transmitted, fluorescence, and reference channels. The

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collected scans for each sample were aligned using a reference yttrium foil and averaged. The

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averaged data were then normalized and the background was removed by spline fitting using the

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XAS data processing program Athena.22 The x-ray absorption near-edge structure (XANES)

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portions of the averaged XAS spectra were used to compare characteristics of the sample and

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reference material spectra. The XANES spectra were also analyzed by the linear combination


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fitting (LCF) data analysis procedure within Athena. This procedure estimated the percentages of

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U(VI) and U(IV), and probed the composition of the U(VI) in each sample.22-24 (See Supporting

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Information Figure S2 for standard spectra and example XANES spectra with LCF analyses for

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two samples).

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Imaging analyses were performed to characterize U on roots from the rewetted planted

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mesocosm and the second mesocosm that was never dried (never dried, planted (2)). Between 8

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mg and 65 mg of roots from each mesocosm were preserved in 1.5 mL microcentrifuge tubes in

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2.5% glutaraldehyde immediately after removal from mesocosms and were kept at 4 °C until

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analyses were performed. The glutaraldehyde was added to preserve the cellular structure of the

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roots.25 Before analysis, root subsamples were gently dried with filter paper and were placed

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into aluminum stubs lined with double-sided carbon tape. Loose sediment from bottom of

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microcentrifuge tubes, which had previously been attached to roots, was deposited in a different

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section of the aluminum post. All sample preparation was performed inside an environmentally

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controlled anoxic chamber filled with argon. Samples were transferred to ambient conditions

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prior to Scanning Electron Microscopy (SEM) analysis. Before placing the sample stubs inside

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the SEM stage, a ~5-10 nm carbon coating was applied to the samples to reduce charging during

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analysis. Focus Ion Beam (FIB) -SEM (FEI Quanta 3D FEG) instrumentation was used for

239

imaging and Energy Dispersive Spectroscopy (EDS) data was collected at several areas along

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roots and on mineral assemblages. The SEM instrument was equipped with an Oxford 80 mm2

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Solid State Detector (SSD) and with INCA software for analyzing the spectra and for

242

determining sample elemental composition. The SEM/EDS sample analysis was performed

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using an electron beam with applied voltage of 30 keV and 1-2 nA current. Each spectrum was

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collected for ~100 seconds with ~20% dead time.


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A second imaging method, autoradiography, was used to further characterize U distributions

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on roots from the planted rewetted mesocosm and from the second planted mesocosm that was

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never dried. This method measured the distribution of radioactivity on roots. Since U was the

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only radioactive element added to the mesocosms and the experimental duration was far shorter

249

than the half-life of U, the distribution of radioactivity is correlated with the U distribution.

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Roots were mounted on cardboard and covered in a Mylar film that allows penetration of alpha

251

radiation. The instrument used to perform the autoradiography measurements was a Fujifilm

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BAS-5000 scanner set for 16 bit, 25 µm pixel digitization. Counting and processing were

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performed following the procedure described by Zeissler et al.26

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Total bacteria, Geobacter spp. (Fe reducers), sulfate-reducing δ-Proteobacteria spp.

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(Desulfuromonadaceae, Desulfobulbaceae, and Desulfobacteraceae), Anaeromyxobacteria spp.

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(metal reducers), and Gallionella-like bacteria (Fe oxidizers) on sediments and roots of dissected

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mesocosms were quantified using DNA isolation and quantitative polymerase chain reaction

258

(qPCR) assay. Root surface-associated soil and minerals were collected by washing the roots

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with 1.5 mL TENP buffer (50 mM Tris[pH 8.0], 20 mM disodium EDTA, 100 mM NaCl, 1%

260

[W/V] PVP) in 2 mL centrifuge tubes. Tubes were centrifuged twice at 1000 rpm for 10 minutes

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and the resulting pellets were saved. DNA was extracted from these root-associated pellets and

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from 500 mg sediment samples using the FastDNA® spin kit for soil (MP Biomedicals, USA) as

263

described by the manufacturer. The concentrations were measured using a Nano-drop 2000

264

spectrophotometer (Thermo Scientific, USA).

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Total bacterial abundance was represented by the number of copies of 16S rRNA genes when

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qPCR was performed with primers BACT1369F-PROK1492R and the TaqMan probe1389F.27

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Sulfate-reducing bacteria were enumerated by qPCR using primer set 361F-685R28 and the


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TaqMan probe 1839F29. Geobacter spp. were enumerated by qPCR using primer set 561F-825R

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and TaqMan probe Gbc2.28 Anaeromyxobacter spp. were enumerated by qPCR using primer set

270

60F-461R.30 Gallionella-like Fe-oxidizing bacteria were enumerated by qPCR using primer set

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628F -998R.31 All qPCR reactions were carried out using a StepOnePlus™ Real-Time PCR

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System (Life Technologies, USA). For DNA quantification, each 20 µL qPCR mixture was

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composed of 10 µL of SYBR Premix Ex Taq® II (Takara, Japan), 0.8 µL of 10 µM of each

274

primer, and ~ 10 ng DNA template. Each assay contained a standard produced by serial dilution

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of plasmids containing specific target genes, independent triplicate templates for each soil

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sample, and triplicate no template controls (NTC).

277 278

Results And Discussion

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Uranium Immobilization in Saturated Mesocosms

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All mesocosms immobilized U very effectively prior to the drying treatment. During this 137-

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day period, the concentration of U in the influent was approximately 9.5 µM (Figure 1-A). Over

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the same period, effluent water was sampled on 21 separate days for U concentration analysis.

283

The concentrations of U in the combined effluents from all mesocosms over all sample days

284

prior to drying varied from 0.002 µM to 0.26 µM with an average of 0.057 µM and a standard

285

deviation of 0.079 µM (Figure 1-A). Individual mesocosms’ effluent water was sampled less

286

frequently and U concentrations exiting a single mesocosm never exceeded 0.31 µM. The

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concentration of U in the effluent of the unplanted mesocosm was within the range of the

288

concentrations in effluents from planted mesocosms on six of the nine days when individual

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mesocosms’ effluents were measured. On the other three days, all within the fourth month after

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U addition started, the unplanted mesocosm’s effluent U concentrations exceeded those of the


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other mesocosms and fell between 0.11 µM and 0.13 µM.

The inconsistency and small

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magnitude of these differences provides little information regarding the role, if any, of plants in

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the initial immobilization of U within the mesocosms.

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XANES-LCF results indicate that immobilized U existed mostly as U(VI) rather than U(IV) in

295

sediments and on roots in all mesocosms. U(IV) accounted for between 4% and 11% of U on

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roots and between 3% and 14% of U in sediments (Table 1). This limited amount of U(IV)

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indicates that bioreduction was not the primary means of U immobilization in the mesocosms

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(Figure 1A). Fe analysis indicates that reducing conditions were present in the sediments of the

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mesocosms that were not dried, where Fe(II) accounted for over 90% of Fe present in 8 out of 10

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sediment samples (see Supporting Information Figure S4). After the drying period, sediments

301

were more oxidized and Fe(II) accounted for, on average, 55% of Fe present in the dried

302

sediments.

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immobilization primarily as U(VI) is consistent with other recent studies of U immobilization in

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model wetlands.11, 13 The strong immobilization of U(VI) would likely not occur in waters with a

305

much higher alkalinity where much of the U(VI) would be complexed as uranyl carbonate which

306

is much more mobile. There was no significant difference between the percent U(IV) in the

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sediments of the unplanted mesocosm and the planted mesocosm which both experienced drying

308

and rewetting. The absence of a plant-related distinction indicates that plant-associated U-

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reducing microbial communities were not solely responsible for the immobilization of U in these

310

mesocosms.

The dominance of U(VI) even in reducing conditions is surprising, but

311

The high level of U(VI) immobilization in these anaerobic wetland mesocosms supports

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observations from previous work in U-contaminated wetland sediments. Much recent work has

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demystified details of controlling variables in reductive immobilization of U(IV). For example,


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314

the U in studies of contaminated French wetlands has been found to be primarily U(IV), its

315

speciation and mobility dependent upon the local concentrations of organic matter and Fe.32, 33

316

In these wetlands, U(IV) was identified both in sediments and in pore water.33 While the current

317

study did not examine the oxidation state of the U in the effluent, the predominance of U(VI) in

318

mesocosm sediments suggests that U(IV) would be minimally or not at all present in mesocosm

319

effluents. In a previous study of microcosms fed with U solution, U(IV) was measured to make

320

up between 14% and 41% of U immobilized on root surfaces.13 This previous study found more

321

U immobilized as U(IV) than the current study did and attributed the presence of U(IV) to U

322

reduction by Geobacter spp. Nonetheless, more than half of immobilized U on root surfaces in

323

both studies was U(VI) which was likely immobilized by abiotic processes such as sorption to

324

root-associated Fe-oxide plaques.13 The possibility of immobile U(VI) under oxic conditions has

325

also been previously observed. For example, U(VI) association with natural organic matter was

326

responsible for high levels of U immobilization in recent experiments with oxic sediments from

327

the Savannah River Site.11 This mechanism could be responsible for the high retention of U(VI)

328

seen in the mesocosms after drying in the current study.

329

Quantitative analyses of bacterial communities also suggest that the majority of the U

330

immobilization was not microbially-mediated.

All mesocosms contained metal-reducing

331

bacteria, with Geobacter spp. generally being most common in sediments and sulfate-reducing

332

bacteria generally being most common on root surfaces (see Supporting Information Tables S2

333

and S3). Geobacter spp. in sediments ranged from 0.0% to 13.5% of the bacteria present, with

334

an average of 1.8%. Sulfate-reducing bacteria on root surfaces ranged from 0.1% to 56.0% of

335

the bacteria present, with an average of 16.6%. Since these metal-reducing bacteria are capable

336

of U reduction, the dominance of immobilized U(VI) over U(IV) indicates that the geochemical


15


Page 16 of 29

337

characteristics of the sediments drove U immobilization more than bioreduction did. Similarly,

338

the number of total bacteria in the 7-11 cm deep bulk sediments of each of the four planted

339

mesocosms was at least 10 times lower than the number of total bacteria in the 4-7 cm deep

340

rhizosphere sediments inside the mesh bags (see Supporting Information Table S2).

341

concentrations of U immobilized in these two sediment horizons (Figure 2-A) did not mirror the

342

disparity in the bacterial populations, suggesting that sorption rather than bacterial processes was

343

primarily responsible for immobilization. Furthermore, SEM/EDS measurements of roots of a

344

mesocosm that was not dried indicated that the U associated with roots was captured in root-

345

associated mineral precipitates (see Supporting Information Figure S3 and Tables S6 – S7-B for

346

elemental composition of these minerals).

347

Effect of Drying on Uranium Mobility

348

The dry period did not cause a large release of U from the mesocosms. Post-drying effluents

349

from both mesocosms briefly contained concentrations of U much higher than those in the pre-

350

drying effluents, but these concentrations were only about 0.27% and 0.08% of the influent U

351

concentration prior to the drying step in the unplanted and planted mesocosms, respectively

352

(Figure 1-B).

353

contained 2.58 µM U and 0.77 µM U, respectively. While these concentrations exceeded the

354

average effluent U concentration from the month prior to drying (0.17 µM U) by factors of 15.6

355

and 4.6, respectively, they quickly approached this pre-drying concentration (Figure 1-B). By

356

the eleventh day after re-saturation, the effluent U concentrations had stabilized around 0.29 µM

357

U and 0.18 µM U in the unplanted and planted mesocosms, respectively. Although the U

358

concentrations in both the unplanted and planted rewetted mesocosms’ effluents exceeded the

359

pre-drying concentrations, these increases were small in magnitude and short-lived. Of the U

The

The first post-drying effluents from the unplanted and planted mesocosms


16

Page 17 of 29


360

estimated to have been immobilized in the mesocosms during the pre-drying period only an

361

estimated 0.45% and 0.29% was released from the unplanted and planted mesocosms,

362

respectively, during the course of the rewetting period.

363

It is possible that the small releases of U after drying and rewetting are related to changes in

364

the oxidation state of the U in the mesocosms. Comparison of all XANES-LCF measurements of

365

sediments from planted mesocosms demonstrated that the percent of U(IV) was generally

366

slightly lower in sediments that had been dried. Although the change in the average percent

367

U(IV) was small enough to possibly be within the noise of the data, dropping from

368

approximately 7% in continuously saturated sediments to approximately 4% in dried or dried and

369

rewetted sediments, Fe measurements indicate it is likely that U(IV) decreased after drying. The

370

ratio of Fe(II) to total Fe decreased in sediments after the 9-day dry period from an average of

371

94% in never dried sediments to an average of 56% in dried sediments (see Supporting

372

Information Figure S4-A), indicating that sediments were less reduced and that oxidation of

373

U(IV) was favorable in sediments during this time. Even after exposure to oxygen, some Fe(II)

374

and U(IV) remained in sediments which is consistent with previous work that found that only a

375

portion of U(IV) in sediments was very reactive with oxygen.10 On root surfaces, the ratio of

376

Fe(II) to total Fe did not change significantly after drying, possibly due to the presence of more

377

oxygen in the root zones throughout the experiment (see Supporting Information Figure S4-B).

378

Accordingly, drying did not seem to alter the percent of U(IV) on the root surfaces. Reoxidation

379

of small amounts of sediment-deposited U(IV) to soluble U(VI) could explain the origins of the

380

U remobilized in effluent following the dry period.

381

It is notable that the initial U pulse after drying was 3.4 times greater from the unplanted

382

mesocosm than from the planted mesocosm (Figure 1-B). This plant-related difference was also


17


Page 18 of 29

383

evident in the pore water that was drained out of the mesocosms when the drying began. The

384

drained pore water from the unplanted mesocosm contained 0.61 µM U which exceeded the

385

average concentration of U in the drained pore water from the planted mesocosms by a factor of

386

15.5. Since the mesocosms contained approximately the same amount of total immobilized U,

387

this slight protection against U remobilization is likely attributable to the biogeochemical activity

388

in the plant’s rhizosphere.

389

Uranium Redistribution towards Roots Following the Dry Period

390

Measurement of U associated with sediments and roots in dissected mesocosms indicated that

391

the dry period caused a localized redistribution of U. The mesocosms that were not dried had

392

more U in the 4-7 cm deep rhizosphere sediment inside the bags than in the 7-11 cm deep bulk

393

sediments (Figure 2-A). In contrast, most of the U in the sediments of the mesocosms that were

394

dried was found in the 7-11 cm deep bulk sediments (Figure 2-A). Both the unplanted and

395

planted rewetted mesocosms held more U in their 7-11 cm deep bulk sediments than in their 4-7

396

cm deep sediments inside the mesh bags, but this downward shift was most pronounced in the

397

unplanted mesocosm whose mesh bag did not contain plant roots (Figure 2-A). These results

398

suggest that drying causes a remobilization and redistribution of U within the mesocosm that is

399

linked to the hydrology of the draining and the absence of plants.

400

Interestingly, this drying-induced movement of immobilized U clarified the importance of

401

roots in protection against net U remobilization. The mesocosms that were dried had at least

402

four times more U on their shallow and deep root surfaces than did the mesocosms that were

403

never dried (Figure 2-B). The source for this additional U on root surfaces is likely some of the

404

U that was dislodged from the 4-7 cm deep rhizosphere sediments inside the mesh bags during

405

draining and then captured by root surfaces.


18

Page 19 of 29


406

Imaging results demonstrate that drying and rewetting change the nature of the U on root

407

surfaces. SEM/EDS analyses of roots that were never dried found U in root-associated minerals

408

(see Supporting Information Figure S3 and Tables S6-S7-B). When the same technique was

409

applied to roots that had been dried and rewetted, U concentrations were below the technique’s

410

detection limit on the deep roots which held far more U than never dried roots according to

411

extraction measurements (see Supporting Information Table S5). This result indicates that U

412

associated with mineral particles on the root surfaces was removed or altered by the drying and

413

rewetting.

414

Analysis of mineral compositions of all spots measured by SEM/EDS indicated that U

415

presence and amount were correlated with phosphorous (P). In spots that contained U, there was

416

also a correlation between amounts of U and sulfur (S). (Supporting Information Figure S5

417

displays these correlations.) The association of immobilized U and P has been documented

418

previously and could be due to co-precipitation.13, 34 Changes in the spatial distributions of S and

419

P during drying and rewetting may help explain the redistribution of U on deep roots during this

420

process. Some of the variability observed could be due to the size of the electron beam that is

421

likely larger (several microns) than individual particles. Therefore coexistence of elements may

422

also be the result of elements detected on different particles.

423

Seventeen spots on deep roots that were not dried were analyzed and eleven spots on deep

424

roots that were dried and rewetted were analyzed. The amount of S in mineral spots analyzed on

425

deep roots differed noticeably after drying and rewetting, increasing from an average of 0.05

426

atomic % before drying to an average of 2.59 atomic % after drying and rewetting. The amount

427

of P on deep roots also increased, but less, from an average of 0.08 atomic % to an average of

428

0.27 atomic %.

While the average masses of S and P seem to increase with drying and


19


Page 20 of 29

429

rewetting, their variabilities do as well. The variability in amounts of S and P in spots analyzed

430

on deep roots, quantified as the standard deviation, changed with drying and rewetting from

431

0.06% to 3.70% and from 0.17% to 0.54% for S and P, respectively. This difference indicates

432

that a change in the composition of some mineral phases could possibly take place during drying

433

and rewetting.

434

Because SEM/EDS could not confirm the high levels of U that had been measured in

435

extractions of dried and rewetted roots in the deep (4-7 cm depth) rhizosphere, autoradiography

436

was performed to understand the U spatial distribution. The highly sensitive autoradiography

437

analyses showed that the entire root sample from the 4-7 cm deep dried and rewetted mesocosm

438

section was white, indicating that radioactivity, and thus U, was located diffusely across the

439

surfaces of roots that had been dried and rewetted (Figure 3). Together, these two sets of data

440

reveal that root-associated U may be immobilized in minerals in a continuously saturated

441

wetland system similar to these mesocosms, but that the drying and rewetting moves U

442

previously immobilized in the rhizosphere sediments towards roots where it binds to the roots’

443

surfaces in a more diffused manner than in the undried system. In sediments without roots, it is

444

possible, and results indicate, that this root-bound U would be mobilized and removed from the

445

system by drying and rewetting. These results build on the previous finding that plant roots’ Fe

446

plaques can harbor U under reducing conditions13, suggesting that plant-associated U could be

447

resilient over seasons to fluctuations in water levels.

448

The importance of plants in protecting against U remobilization during drying is confirmed by

449

measurements of the concentrations of U in drained pore water. As mentioned above, the

450

drained pore water from the unplanted mesocosm contained 15.5 times more U than did the

451

drained pore water from the planted mesocosms. U appears to sorb directly on roots and may


20

Page 21 of 29


452

sorb or co-precipitate with Fe(III) onto iron plaques that develop on wetland plant root surfaces,

453

thereby immobilizing U(VI) that might result from the oxidation of U(IV) during periods when

454

sediments become oxic. These results demonstrate that the presence of plants may lower U

455

mobility during water table fluctuations in contaminated wetlands.

456 457

ASSOCIATED CONTENT

458

Supporting Information

459

Data from qPCR analysis of mesocosms, sample XANES spectra with LCF analyses, XANES

460

standard spectra, calculated speciation of U in influent mixture, sample SEM/EDS data from a

461

never dried root sample, elemental composition of root-associated minerals measured by

462

SEM/EDS, plots of SEM/EDS U, P, and S data to show correlation, and measurements of Fe(II)

463

in mesocosm sediments and roots are included in Supporting Information. This material is

464

available free of charge via the Internet at http://pubs.acs.org.

465

AUTHOR INFORMATION

466

Corresponding Author

467

*Address: Princeton University, Department of Civil and Environmental Engineering, E-Quad,

468

Princeton, NJ 08540; Phone: (609) 258-4653; Email: [email protected].

469

Present Addresses

470

†Biotechnology Development and Applications, CB&I Federal Services, 17 Princess Rd,

471

Lawrenceville, New Jersey 08648, United States.

472

Notes


21


Page 22 of 29

473

The authors declare no competing financial interests.

474

ACKNOWLEDGMENTS

475

This research was supported through contract DE-SC0006847 by the Subsurface

476

Biogeochemical Research Program of the U.S. Department of Energy's Office of Biological and

477

Environmental Research.

478

A portion of this research used resources of the Advanced Photon Source, a U.S. Department

479

of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by

480

Argonne National Laboratory under Contract No. DE-AC02-06CH11357. The Materials

481

Research Collaborative Access Team (MRCAT) operations at the Advanced Photon Source are

482

supported by the DOE and the MRCAT member institutions. Although the U.S. Environmental

483

Protection Agency (EPA) contributed to some of the work described in this document, the

484

research presented was not performed by or funded by EPA and was not subject to EPA's quality

485

system requirements. Consequently, the views, interpretations, and conclusions expressed in this

486

document are solely those of the authors and do not necessarily reflect or represent EPA's views

487

or policies.

488

A portion of this research was performed using Radiochemistry Annex at EMSL, a DOE

489

Office of Science User Facility sponsored by the Office of Biological and Environmental

490

Research and located at Pacific Northwest National Laboratory.

491

REFERENCES

492 493 494 495 496

1. Arnold, C. The legacy of uranium on the Navajo Nation once upon a mine. Environ. Health Persp. 2014, 122 (2), A44-A49. 2. Dias da Cunha, K. M.; Henderson, H.; Thomson, B. M.; Hecht, A. A. Ground water contamination with (238)U, (234)U, (235)U, (226)Ra and (210)Pb from past uranium mining: Cove Wash, Arizona. Environ. Geochem. Health 2014, 36 (3), 477-87.


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3. Kaplan, D. I.; Serkiz, S. M. Quantification of thorium and uranium sorption to contaminated sediments. J. Radioanal. Nucl. Ch. 2001, 248 (3), 529-535. 4. Kurttio, P.; Auvinen, A.; Salonen, L.; Saha, H.; Pekkanen, J.; Makelainen, I.; Vaisanen, S. B.; Penttila, I. M.; Komulainen, H. Renal effects of uranium in drinking water. Environ. Health Persp. 2002, 110 (4), 337-342. 5. Kurttio, P.; Komulainen, H.; Leino, A.; Salonen, L.; Auvinen, A.; Saha, H. Bone as a possible target of chemical toxicity of natural uranium in drinking water. Environ. Health. Persp. 2005, 113 (1), 68-72. 6. Leggett, R. W.; Pellmar, T. C. The biokinetics of uranium migrating from embedded DU fragments. J. Environ. Radioactiv. 2003, 64 (2-3), 205-225. 7. Lovley, D. R.; Phillips, E. J. P.; Gorby, Y. A.; Landa, E. R. Microbial Reduction of Uranium. Nature 1991, 350 (6317), 413-416. 8. Groudev, S.; Spasova, I.; Nicolova, M.; Georgiev, P. In situ bioremediation of contaminated soils in uranium deposits. Hydrometallurgy 2010, 104 (3-4), 518-523. 9. Senko, J. M.; Istok, J. D.; Suflita, J. M.; Krumholz, L. R. In-situ evidence for uranium immobilization and remobilization. Environ. Sci. Technol. 2002, 36 (7), 1491-1496. 10. Sharp, J. O.; Lezama-Pacheco, J. S.; Schofield, E. J.; Junier, P.; Ulrich, K. U.; Chinni, S.; Veeramani, H.; Margot-Roquier, C.; Webb, S. M.; Tebo, B. M.; Giammar, D. E.; Bargar, J. R.; Bernier-Latmani, R. Uranium speciation and stability after reductive immobilization in aquifer sediments. Geochim. Cosmochim. Ac. 2011, 75 (21), 6497-6510. 11. Li, D.; Seaman, J. C.; Chang, H. S.; Jaffe, P. R.; van Groos, P. K.; Jiang, D. T.; Chen, N.; Lin, J. R.; Arthur, Z.; Pan, Y. M.; Scheckel, K. G.; Newville, M.; Lanzirotti, A.; Kaplan, D. I. Retention and chemical speciation of uranium in an oxidized wetland sediment from the Savannah River Site. J. Environ. Radioactiv. 2014, 131, 40-46. 12. Wan, J. M.; Dong, W. M.; Tokunaga, T. K. Method to Attenuate U(VI) Mobility in Acidic Waste Plumes Using Humic Acids. Environ. Sci. Technol. 2011, 45 (6), 2331-2337. 13. Chang, H. S.; Buettner, S. W.; Seaman, J. C.; Jaffe, P. R.; Koster van Groos, P. G.; Li, D.; Peacock, A. D.; Scheckel, K. G.; Kaplan, D. I. Uranium immobilization in an iron-rich rhizosphere of a native wetland plant from the Savannah River Site under reducing conditions. Environ. Sci. Technol. 2014, 48 (16), 9270-9278. 14. Sandino, A.; Bruno, J. The solubility of (UO2)3(PO4)2.4H2O(S) and the formation of U(VI) phosphate complexes - their influence in uranium speciation in natural-waters. Geochim. Cosmochim. Ac. 1992, 56 (12), 4135-4145. 15. Moon, H. S.; Komlos, J.; Jaffe, P. R. Uranium reoxidation in previously bioreduced sediment by dissolved oxygen and nitrate. Environ. Sci. Technol. 2007, 41 (13), 4587-4592. 16. Singh, G.; Sengor, S. S.; Bhalla, A.; Kumar, S.; De, J.; Stewart, B.; Spycher, N.; Ginn, T. M.; Peyton, B. M.; Sani, R. K. Reoxidation of biogenic reduced uranium: a challenge toward bioremediation. Crit. Rev. Env. Sci. Tec. 2014, 44 (4), 391-415. 17. Dong, W.; Wan, J. Additive surface complexation modeling of uranium(VI) adsorption onto quartz-sand dominated sediments. Environ. Sci. Technol. 2014, 48 (12), 6569-6577. 18. Hoagland, D. R.; Arnon, D. I. The water-culture method for growing plants without soil; College of Agriculture, University of California: Berkeley, CA, 1950; Vol. 347. 19. Komlos, J.; Peacock, A. A.; Kukkadapu, R. K.; Jaffe, P. R. Long-term dynamics of uranium reduction/reoxidation under low sulfate conditions. Geochim. Cosmochim. Ac. 2008, 72, 3603-3615.


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20. Komlos, J.; Jaffe, P. R. Effect of iron bioavailability on dissolved hydrogen concentrations during microbial iron reduction. Biodegradation 2004, 15 (5), 315-325. 21. Segre, C. U.; Leyarovska, N. E.; Chapman, L. D.; Lavender, W. M.; Plag, P. W.; King, A. S.; Kropf, A. J.; Bunker, B. A.; Kemner, K. M.; Dutta, P.; Duran, R. S.; Kaduk, J. In The MRCAT Insertion Device Beamline at the Advanced Photon Source, Synchrotron Radiation Instrumentation: Eleventh U.S. National Conference, New York, 2000; Pianetta, P., Ed. American Institute of Physics: New York, 2000; pp 419-422. 22. Ravel, B.; Newville, M. ATHENA, ARTEMIS, HEPHAESTUS: data analysis for X-ray absorption spectroscopy using IFEFFIT. J. Synchrotron. Radiat. 2005, 12, 537-541. 23. Bunker, G. Introduction to XAFS a practical guide to X-ray absorption fine structure spectroscopy. In Cambridge University Press: Cambridge, UK, 2010. 24. Kelly, S. D.; Hesterberg, D.; Ravel, B. Analysis of Soils and Minerals Using X-ray Absorption Spectroscopy. In Methods of Soil Analysis, Part 5 - Mineralogical Methods, Ulery, A. L.; Drees, L. R., Eds. Soil Science Society of America: Madison, WI, USA, 2008; p 367. 25. Chrispeels, M. J.; Vatter, A. E. Preservation of Ultrastructure of Plant Cells Using Glutaric Acid Dialdehyde as a Fixative. Nature 1963, 200 (490), 711. 26. Zeissler, C. J.; Lindstrom, R. M.; McKinley, J. P. Radioactive particle analysis by digital autoradiography. J. Radioanal. Nucl. Ch. 2001, 248 (2), 407-412. 27. Suzuki, M. T.; Taylor, L. T.; DeLong, E. F. Quantitative analysis of small-subunit rRNA genes in mixed microbial populations via 5 '-nuclease assays. Appl. Environ. Microb. 2000, 66 (11), 4605-4614. 28. Stults, J. R.; Snoeyenbos-West, O.; Methe, B.; Lovley, D. R.; Chandler, D. P. Application of the 5 ' fluorogenic exonuclease assay (TaqMan) for quantitative ribosomal DNA and rRNA analysis in sediments. Appl. Environ. Microb. 2001, 67 (6), 2781-2789. 29. N'Guessan, A. L.; Moon, H. S.; Peacock, A. D.; Tan, H.; Sinha, M.; Long, P. E.; Jaffe, P. R. Postbiostimulation microbial community structure changes that control the reoxidation of uranium. Fems. Microbiol. Ecol. 2010, 74 (1), 184-195. 30. Petrie, L.; North, N. N.; Dollhopf, S. L.; Balkwill, D. L.; Kostka, J. E. Enumeration and characterization of iron(III)-reducing microbial communities from acidic subsurface sediments contaminated with uranium(VI). Appl. Environ. Microb. 2003, 69 (12), 7467-7479. 31. Wang, J. J.; Vollrath, S.; Behrends, T.; Bodelier, P. L. E.; Muyzer, G.; Meima-Franke, M.; Den Oudsten, F.; Van Cappellen, P.; Laanbroek, H. J. Distribution and diversity of Gallionella-like neutrophilic iron oxidizers in a tidal freshwater marsh. Appl. Environ. Microb. 2011, 77 (7), 2337-2344. 32. Wang, Y.; Bagnoud, A.; Suvorova, E.; McGivney, E.; Chesaux, L.; Phrommavanh, V.; Descostes, M.; Bernier-Latmani, R. Geochemical Control on Uranium(IV) Mobility in a MiningImpacted Wetland. Environ.l Sci.Technol. 2014, 48, 10062-10070. 33. Wang, Y.; Frutschi, M.; Suvorova, E.; Phrommavanh, V.; Descostes, M.; Osman, A. A. A.; Geipel, G.; Bernier-Latmani, R. Mobile uranium(IV)-bearing colloids in a mining-impacted wetland. Nature Communications 2013, 4. 34. Zhou, P.; Gu, B. H. Extraction of oxidized and reduced forms of uranium from contaminated soils: Effects of carbonate concentration and pH. Environ. Sci. Technol. 2005, 39 (12), 4435-4440.


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Page 25 of 29


U Concentration (µ µM)

A. U in Mesocosm Influent and Effluent Before Drying Period 10 8 6 4 2 0 0

20

40

60 80 Days

100 120 140

Average U in influent before drying Average U in effluent before drying

585

U concentration (µ µM)

B. U in Mesocosm Effluent After Drying and Rewetting 3.0 2.5 2.0 1.5 1.0 0.5 0.0 145

147

149

151 153 155 Days U in unplanted mesocosm effluent

157

U in planted mesocosm effluent

586 587

Average U in effluent in month prior to drying

Figure 1. U in mesocosm influent and effluent before (A) and after (B) drying period.


25


Page 26 of 29

A. U on Mesocosm Sediments 350 300 µg U / g dry soil

250 200 150 100 50 0

Sediment Type and Depth After drying, planted After rewetting, planted After rewetting, unplanted Never dried, planted (1) Never dried, planted (2)

588

µg U / g dry root

B. U on Mesocosm Roots 6000 4000 2000 0

589

5-10 cm 0-5 cm Root Depth After drying After rewetting Never dried (1) Never dried (2)

590

Figure 2. U on mesocosm sediments and root surfaces. In the four planted mesocosms, “In Bag”

591

sediments are rhizosphere sediments. Bars indicate the average of U concentrations of two

592

samples from each location and error bars indicate the samples’ ranges.


26

Page 27 of 29


593 594

Figure 3. Distribution of radioactivity on roots captured by autoradiography. Individual roots

595

(indicated by the dotted lines in the figure) from each location were placed on the circular dark

596

plates in the middle of each panel. White spots on the roots indicate high U concentrations.

597


27


Page 28 of 29

598

Table 1. U(IV) and U(VI) as percentages of total U in sediments and on root surfaces in

599

mesocosms. XANES-LCF measurements to determine the oxidation state of U were made on a

600

single root or sediment sample from each horizon. In a number of cases, U concentrations were

601

under the detection limit (UDL) for the technique.

Depth Bulk Sediments After drying, planted

Rhizosphere Sediments Root Surfaces Bulk Sediments

After rewetting, planted

After rewetting, unplanted

Rhizosphere Sediments Root Surfaces Bulk Sediments Sediments in Bag Bulk Sediments

Never dried, planted (1)

Rhizosphere Sediments Root Surfaces Bulk Sediments

Never dried, planted (2)

Rhizosphere Sediments Root Surfaces

1-4 cm 4-7 cm 7-11 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm 7-11 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm 7-11 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm 7-11 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm 7-11 cm 1-4 cm 4-7 cm 1-4 cm 4-7 cm

% U(IV) of % U(VI) of total U total U 3 97 UDL 3 97 UDL UDL 10 90 11 89 UDL UDL 3 97 5 95 5 95 8 92 7 93 UDL UDL 6 94 4 96 5 95 UDL UDL 7 93 14 86 4 96 8 92 9 91 UDL 3 97 6 94 6 94 7 93 4 96 8 92

602


28

Page 29 of 29


603 604

Table of Contents Graphic


29

Uranium Redistribution Due to Water Table Fluctuations in Sandy

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