Effect of Biofouling on the Performance of ... - ACS Publications

Subscriber access provided by NEW YORK UNIV

Article

Effect of biofouling on the performance of amidoxime-based polymeric uranium adsorbents Jiyeon Park, Gary A. Gill, Jonathan E. Strivens, Li-Jung Kuo, Robert Jeters, Andrew Avila, Jordana Wood, Nicholas J Schlafer, Chris J. Janke, Erin A Miller, Mathew Thomas, R. Shane Addleman, and George Bonheyo Ind. Eng. Chem. Res., Just Accepted Manuscript • DOI: 10.1021/acs.iecr.5b03457 • Publication Date (Web): 27 Jan 2016 Downloaded from http://pubs.acs.org on January 28, 2016

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Industrial & Engineering Chemistry Research is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Industrial & Engineering Chemistry Research

Effect Of Biofouling On The Performance Of Amidoxime-Based Polymeric Uranium Adsorbents Jiyeon Parka, Gary A. Gilla, Jonathan E. Strivensa, Li-Jung Kuoa, Robert T. Jetersa, Andrew Avilaa, Jordana R. Wooda, Nicholas J. Schlafera, Christopher J. Jankec, Erin A. Millerb, Mathew Thomasb, R. Shane Addlemanb, and George T. Bonheyoa a. Marine Sciences Laboratory, Pacific Northwest National Laboratory, 1529 W. Sequim Bay Rd, Sequim, WA 98382, United States b. Pacific Northwest National Laboratory, 902 Battelle Boulevard, Richland, WA 99352, United States c. Oak Ridge National Laboratory, P.O. Box 2008, Oak Ridge, TN 37831, United States

ABSTRACT: The Marine Science Laboratory at the Pacific Northwest National Laboratory evaluated the impact of biofouling on uranium adsorbent performance. A surface modified polyethylene adsorbent fiber provided by Oak Ridge National Laboratory, AF adsorbent, was tested either in the presence or absence of light to simulate deployment in shallow or deep marine environments. Samples of the adsorbent fiber were exposed to seawater as loose fibers packed with glass beads in columns and as >10cm long braids of fiber placed in a flume that provided a continuous flow representative of natural ocean currents. 42-day exposure tests in column and flume settings showed that biofouling resulted in decreased uranium uptake by the adsorbent fiber. Uranium uptake was reduced by up to 30 %, in the presence of simulated sunlight, which also increased biomass accumulation and altered the microbial community composition on the fibers. These results suggest that deployment below the photic zone would mitigate the effects of biofouling, resulting in greater yields of uranium extracted from seawater.

INTRODUCTION The Fuel Resources Program at the U.S. Department of Energy’s Office of Nuclear Energy is supporting the development of adsorbent technology to extract uranium from seawater. This technology is being developed to provide a sustainable and economically viable supply of uranium fuel for nuclear reactors 1. Uranium is present in seawater at a concentration of ~3.3 ppb, amounting to a global marine resource of an estimated 4.5 billion metric tons 2. A major effort in the development of this technology at the Pacific Northwest National Laboratory (PNNL) is to test the performance of the uranium adsorption materials in natural seawater under realistic marine conditions. Briefly, the envisioned strategy of the program is to incorporate uranium adsorbent chemistry into a fibrous form (e.g., through grafting onto polyethylene fibers) and to create large (i.e., 10s of meters long) braids of the material that may be anchored to the seafloor during the collection (adsorption) process 1, Preliminary testing and selection of amidoxime-based adsorbent fibers was performed at a number of participating laboratories using artificial seawater that lacked many of the chemical and biological properties of natural seawater. Additional evaluations were then performed at the PNNL Marine Science Laboratory (MSL) in Sequim, WA using filtered seawater. Although artificial and filtered seawater was sufficient for an initial down-selection of both support materials and ligand chem-

istry, the lack of cellular, biomolecular and organic matter in these test waters was unrealistic. Design features of the adsorbent materials, such as large porous or textured surface areas with micro or nano-scaled features that are intended to maximize uranium uptake may also enhance the adsorption of dissolved and particulate organic matter and colonization by marine microorganisms 3-6. Biofouling is the accumulation of microorganisms, algae, plants or animals on wetted surfaces. In the marine environment, biofouling is generally a four-step process 7, 8. During the first stage, surfaces are rapidly coated with an organic conditioning film beginning within 5 to 10 seconds after immersion 9. Then single bacterial cells and diatoms begin to settle, adhere and colonize on the surface 10. During the third stage, microbial films develop creating rough surfaces that trap more particles and organisms including the larval forms of macroorganisms such as barnacles. During the final stage, outgrowth of macroorganisms like mussels, barnacles or algae occurs on the fouled surface 11, 12. In marine industries, biofouling causes various problems including increased shipping costs due to ship hull fouling (greater fuel consumption), increased maintenance cost and time, the release of toxins in antifouling paints during cleaning, and the unintended transport of invasive foreign species 13-17.

1 ACS Paragon Plus Environment

Industrial & Engineering Chemistry Research Concerns about the impact of biofouling on uranium adsorbent performance may be summarized as shown in Figure 1. In the absence of biofouling, the amount of uranium uptake should increase until the adsorbent is saturated as shown by the blue curve. However, prolonged deployment of uranium adsorbent in the ocean will increase exposure to live and dead biogenic materials and allow for the growth of any cells that colonize the surface of the adsorbent material (green curve). An increase in biomass will have two primary effects that decrease uranium uptake and recovery (red curve): 1) limiting the accessibility of the ligands to ocean water during uptake and 2) interfering with the uranium recovery process by diluting the extraction solution and/or restricting its access to the ligands. A third potential detrimental effect that is not covered in this study is that fouling may lead to decreased reusability of the adsorbent due to biocorrosion, detrimental treatments necessary to remove fouling, damage from added weight and drag, or promoting damage by fish and invertebrates grazing on organic matter found on the surface of the adsorbent. Therefore, in order to maximize uranium extraction performance in a real marine environment, it is important to understand how biofouling impacts uranium uptake and recovery, potential rates of buildup, the correlation of biomass to interference, and the kinetics of the adsorbents and their durability. This information will be highly valuable and extensively useful in developing deployment and operation plans and identifying cost effective and compatible methods to mitigate the effects of biofouling.

Environmental Effects (Conceptual Figure)

Biomass accumulation (g biomass/kg adsorbent)

U-adsorption (g U/kg adsorbent)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 13

maximum potential impact fouling might have on the adsorbents. A second set of experiments considered how different deployment strategies, shallow water and light exposed or deep water and no lighting, might affect performance. Sunlight plays an important role in biofouling because in many of the early stage, surface-colonizing microorganisms are photosynthetic (e.g., diatoms, cyanobacteria). In this study, pre-weighed samples of adsorbent material were exposed either to filtered or unfiltered seawater, with or without pre-treatment with microorganisms (to maximize fouling and establish a theoretical upper bound on effects) and with or without lighting to simulate shallow or deep seawater settings. Some experiments were performed using adsorbent material packed into columns to strictly control and measure the amount of water passing over the fibers and to assure that nearly 100 % of the fiber could be recovered at the conclusion of a test. This column set up has been previously described 2. A limitation of the columns was that these cannot be used with unfiltered seawater due to rapid clogging of the filter bed that holds the adsorbent. Therefore, additional experiments were conducted using braids of fiber in an open flume setting with flowing unfiltered seawater, with or without light. A limitation of these flume studies was the potential loss of loose adsorbent material and the ability to accurately measure the mass of adsorbent (excluding biofouling and water) in recovered samples. Measurements quantified the amount of biofouling material and the total amount of uranium adsorbed onto the fiber using a destructive method of analysis. A DNA-based approach was used to assess the composition and diversity of the microbial communities found on the adsorbent. An attempt was also made to visualize fouling on the surface of adsorbent fibers to better understand where fouling occurs relative to the surface structure of the fiber. These data could be used to gain insight into structural designs that might maximize the water accessible surface area of the adsorbent while limiting the extent and impact of biofouling. EXPERIMENTAL METHODS Adsorbents

Exposure Time (Days) U-adsorption per adsorbent (g/kg) without biofouling U-adsorption per adsorbent (g/kg) with biofouling Biomass accumulation per adsorbent (g/kg)

Figure 1. Environmental impact on uranium extraction from seawater. Conceptual figure showing the relationship of time with biomass accumulation (biofouling) and its effect on uranium uptake and recovery.

To quantify the potential impact of fouling on uranium uptake, a set of time series experiments were conducted that examined biomass accumulation, cell growth, and uranium uptake on a representative, leading adsorbent material. One set of experiments was designed to assess the

ORNL developed and produced AF1 and AI8 adsorbents. Adsorbents were prepared using hollow-gearshaped, high surface area polyethylene (PE) fibers and a radiation-induced graft polymerization (RIGP) method 2. AF1 and AI8 adsorbents were different in the grafting comonomer (itaconic acid and vinylphosphonic acid, respectively). Both used amidoxime as the uranium binding ligand. Column system Marine testing was conducted using ambient seawater from Sequim Bay, WA. The PNNL Marine Science Laboratory (MSL) seawater delivery system provided am-


Page 3 of 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


bient seawater into a wet laboratory for scientific investigation. Ambient seawater was drawn continuously (day and night) by pump from a depth of ~10 m from Sequim Bay; consequently water was collected through all diurnal and tidal cycles. Input water is delivered through a plastic pipe and was passed through an Arkal Spin Klin™ filter system (nominal pore size 40 µm) to remove large particles. The seawater was then fed in a large volume outdoor reservoir tank and supplied to the laboratory research facilities at MSL by gravity feed through PVC piping. Seawater from the large reservoir tank was fed sequentially through 5 µm and then 1 µm cellulose filters and collected in a 180 L fiberglass reservoir tank (head tank). Seawater in the head tank could be heated to a desired temperature. Temperature-controlled seawater was drawn from the head tank with a pump (non-metallic pump head), passed through a 0.35 to 0.45 µm polyethersufone (Memtrex MP, GE Power and Water) or cellulo bromoperoxidase membrane cartridge filter and into a 24-port PVC manifold. Water that was not used to expose adsorbent material passed through the manifold and was returned to the head tank. Pressure in the manifold was controlled with a gate valve at the outlet of the manifold. MSL has four separate 24-port manifolds, linked to three separate head tanks, which permitted testing of 96 adsorbent materials in flow-through columns simultaneously (Fig. 2).

Figure 2. Column experiment set-up either in the absence or presence of light. (A) Columns were covered with aluminum foil to prevent exposure to light. (B) Daylight spectrum light was provided on a 12:12 hour light-dark cycle. A fan was placed to prevent heating of the columns.

impact of biofouling on uranium uptake, particularly in the column experiments where minimal cellular material was expected to be present in the filtered seawater. Column experiment, either in the presence or absence of light Pre-weighed ORNL AF1 adsorbent was conditioned in 2.5 % KOH at 80 °C for 1 hour. Conditioned adsorbents with or without pre-fouling were placed between packing materials (acid cleaned 5 mm glass beads and glass fiber) in columns. Columns were mounted on manifolds and 0.45 µm filtered seawater was passed through these for 42 days at 20 °C. To prevent exposure to light, columns were wrapped in aluminum foil (Fig. 2A). For column testing with light, daylight spectrum light (5000K) was provided on a 12:12 hour light-dark cycle and a fan was used to prevent overheating of the columns (Fig. 2B). Flume System Ambient, unfiltered seawater was drawn by pump from a depth of ~10 m from Sequim Bay through a plastic pipe, and delivered through a PVC piping system to the wet laboratory for use. Water was drawn continuously through all diurnal and tidal cycles. Furthermore, the continuous operation of the pumping system over several months prior to the experiments afforded species from multiple seasons and all diurnal and tidal cycles with an opportunity to colonize the plumbing and tanks, thus providing a potential secondary source of organisms. Gross filtration of the ambient seawater was conducted to remove large debris using a Big Bubba® non-metallic filter housing and a 150 µm filter. The 150-µm filtration allowed for the free passage of most phytoplankton species that contribute to biofouling, but removed larger marine plankton species. Temperature control for experimental purposes was achieved by feeding the 150 µm filtered seawater into a 180 L head tank that had a 10,000 W titanium immersion heater. The ambient temperature of seawater as it entered the head tank was ~10 o C and the temperature in the head tank was maintained at 20 °C, with a variability of approximately ± 1.5 °C. Temperature controlled water was drawn out of the head tank using a pump and delivered to a multi-port manifold on the wetlab table for experimental distribution.

Pre-fouling with Navicula incerta A stock culture of the marine benthic diatom Navicula incerta (UTEX 2044) was grown in F/2 medium (NCMA, ME) using conditions described below. ORNL AF1 adsorbent was weighed (50 mg) and then conditioned in 2.5 % KOH at 80 °C for 1 hour. Conditioned fibers were pH neutralized and then placed in a 50 mL plastic vial and submerged in deionized water to remove most of the KOH solution. Fibers were transferred to sterile 250 mL glass flasks each containing 150 mL (6.7 x 104 cells/mL) of the N. incerta culture. Fibers and culture mixtures were then incubated at room temperature with shaking (135 rpm) and daylight spectrum light (5000K) on a 12:12 hour light-dark cycle until fouling became visible. Pre-fouling was designed to ensure biological growth and test the maximum

PNNL developed recirculating flumes for conducting exposure tests with braided adsorbent materials under controlled temperature and flow-rate conditions. For exposures with 150 µm filtered seawater, the flumes were constructed of an acrylic material to allow ambient light to pass to the seawater in the flume. Allowing light penetration permitted the growth of photosynthetic organisms (biofouling) as would occur in the natural photic seawater environment. Two translucent flumes were built for the biofouling study. Flume experiment The braided ORNL AI8 adsorbent was tested in clear flumes either in the presence or absence of light. Di-



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

urnal daylight spectrum light (5000K) was provided for the light flume, and the dark flume was covered with a black tarp to prevent exposure to light (Fig. 3). The intensity of light at the surface of the water in the flume was measured to be approximately 50 µmol of photons m-2 sec -1, a small fraction of the light intensity of a bright summer day at noon (~ 2,000 µmol of photons m-2 sec-1), yet still strong enough to enhance photosynthetic growth inside the flume as evidenced by recurring visible algae growth on the flume walls.

Page 4 of 13

different time points; Time 0, 7 days, 14 days, 21 days, 28 days and 42 days.

Figure 4. ORNL AI8 adsorbent. (A) Mini braids were preweighed (100 mg) and tied using fishing line prior to KOH conditioning. (B) Conditioned mini braids were secured inside the flume using cable ties. Red box indicates a single mini braid. (C) Braided AI8 before conditioning. (D) Conditioned AI8 braid was secured inside the flume.

Uranium and trace element analysis

Figure 3. Flume set up. In order to test the impact of light on biofouling and subsequent uranium uptake, two clear flumes, Dark Flume and Light Flume, were set up. 150 µm filtered seawater (to remove large particles) was drawn to the flumes by pumps at 2~5 L/min.

The adsorbent braids were tested in two formats: ‘regular’ braids (9 g) and pre-measured ‘mini-braids’ (100 mg) that were prepared as follows. A braid of adsorbent material was made by ORNL. This braid consisted of long (10s of meters) fibers that were braided together to form a tight central stem surrounded by loose loops of fiber in which the each loop was 6 to 10 cm in circumference. Shorter sections were cut at the stem from the primary preparation to create ~10-15 cm long (9g) sections of braided material herein referred to as ‘regular braids.’ Mini braids were made by cutting loops off of the stem to create clusters of now separate, linearized fiber pieces. The minibraids were weighed and subsequently tied mid-length using a fishing line prior to KOH conditioning (Fig. 4A). Prepared braids (which had also been KOH conditioned) were secured inside the flume using cable ties and exposed to the coarsely filtered (150 µm) seawater at 20 °C (Fig. 4B & 4D). The regular braids provided a format intended for actual deployment in the environment, but fouling on the surface appeared to be uneven, the size of the sample was deemed large enough to pose a radiation safety challenge, and smaller samples cut from the regular braid after exposure could not be weighed accurately as these now contained seawater, biofouling, and accumulated metals. The pre-weighed, smaller mini-braids provided greater accuracy for measurements dependent upon the starting mass of material, but may have had introduced some differences due to the altered physical format. Samples were collected at 6

Following exposure to seawater, adsorbent samples were rinsed in DI water to remove accumulated salts. Each sample was then placed in a plastic vial and dried using a heating block (80 °C overnight). The dried and weighed adsorbent sample was placed into a plastic or Teflon container and then a solution of 50 % aqua regia was added for a total digestion of the adsorbent. The digestion mix was incubated in a heating block at 80 °C for 2 hours, and then analyzed for uranium and trace elements via ICPOES. Adsorption (uptake) was determined based on the mass of the recovered elements per mass of adsorbent (g of element adsorbed per kg of dry adsorbent). Microscopy Adsorbent fiber samples were examined under a Leica DMIRB Inverted Fluorescence Microscope (Bartels & Stout, Inc., WA) at 1000 X magnification. Non-purgeable organic carbon analysis The adsorbent samples (50 mg) were placed in a 50 mL centrifuge tube to which 30 mL of 3 % hydrogen peroxide solution was added and then vortexed for 10 seconds to strip the accumulated biomass off of the adsorbent fiber. The 3 % hydrogen peroxide solution was made by diluting 30 % hydrogen peroxide (JT Baker, PA) with 18MΩ Milli-Q water (1:10 ratio). The tube was then placed in a 55 °C water bath. After incubating for 30 minutes, the tube was removed and vortexed for an additional 10 seconds, then placed back into the water bath for another 30minute incubation. Following these incubations, the 50 mL conical tube containing fiber sample was fastened to the ultrasound horn of a VialTweeter ultrasound unit (Hielscher USA, Inc., NJ) by two large rubber bands and aligned such that fiber sample inside was directly facing the end of the horn. Sonication was performed for one minute (100 % amplitude, 0.75 cycle) and then the tube was rotated 180 degrees and sonication was repeated to ensure thorough treatment of the fiber surface. Non-purgeable organic carbon (NPOC) analysis was performed using a Shimadzu TOC-L (Shimadzu Scien-


Page 5 of 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


tific Instruments, Inc., MD). A 30 mL sample of the hydrogen peroxide solution used to clean fiber samples was placed in a 40 mL certified clean NPOC test vial (Fisher Scientific, NH) for NPOC testing. Potassium hydrogen phthalate (50 ppm and 5 ppm) and unused 3 % hydrogen peroxide were used as test controls. All NPOC vials were loaded into the Shimadzu ASI-L auto sampler. NPOC settings were as follows: Sparge time 2 minutes, acid addition 1.5 % HCl trace grade (Fisher Scientific, PA), 3 of 5 injections per sample, C.V. Max 2 %. To subtract any carbon input from the solvents, samples of the 3% hydrogen peroxide solution and Milli-Q water were taken and tested in the NPOC analysis as blank controls. DNA extraction and community analysis A 50 mL centrifuge tube containing fiber sample was filled with 25 mL nuclease free water (Bioexpress, UT), and vortexed to release biomass accumulation from the fiber into the water. The cell suspension, excluding adsorbent fibers, was carefully transferred by pipette into a sterile 50 mL centrifuge tube and then further split into two equivalent fractions. The fractions were centrifuged at 4150 rpm for 30 minutes at 4 °C. The supernatant of each fraction was decanted, resulting in two pellets. One pellet was resuspended in 480 µL nuclease free water and placed in a sterile 2 mL microcentrifuge tube to be used for direct PCR amplification. The other pellet was resuspended in 480 µL 50 mM EDTA and then transferred to a sterile 2 mL microcentrifuge tube to be further processed for total DNA extraction using the Wizard Genomic DNA Purification Kit (Promega, WI). DNA amplification of the bacterial 16S23S internal transcribed spacer (ITS) region and ITS15.8S0ITS2 region of eukaryotes was performed. Polymerase chain reaction (PCR) was performed using OneTaq DNA polymerase (New England Biolabs, MA). Template for the PCR reaction consisted of either purified DNA from the total DNA extraction process or raw suspension for direct amplification. The PCR products were run on a Labchip GXII bioanalyzer (Perkin Elmer, MA) using the High Sensitivity Lab-Chip and Reagent Kit in order to determine amplicon sizes and relative amounts of the amplified DNA.

% stock solution diluted with 1 X PBS) for 5 minutes at room temperature followed by washing twice with 1X PBS for two times. A control fiber sample that was only exposed to sterile culture medium was also prepared for imaging. The wet fibers were then shipped to the Advanced Photon Source (APS) at Argonne National Laboratory in sterile seawater media. X-ray microtomography was performed at the Advanced Photon Source, beamline 2-BM 20. The dry fibers were transferred into a plastic pipette tip, which provided stability with low attenuation. The fibers were imaged at a resolution of 0.7 µm per pixel; a series of images are acquired at different sample angles over the course of 5-10 minutes to form a 3-D dataset, which is then reconstructed into a volume of 2048 x 2048 x 2048 pixels. The x-ray beam energy was 22.5 keV, with the detector 200 mm back from the sample. This distance allowed for edge enhancement due to refractive index effects, which increases contrast on low-atomic number materials. After reconstruction into a volume, the dataset was then rendered into a 2-D image using Matlab. The color scale in the image indicates the relative absence of intensity, either due to attenuation or refractive effects, and tends to be larger for higher atomic number materials or near large changes in electron density. RESULTS AND DISCUSSION Effect of biofouling on uranium uptake in column testing

Advanced photon source (APS) imaging

The objective of the seawater uranium extraction program in the United States, Japan, and China is to develop materials and an economically viable process that can compete with traditional terrestrial mining resources 2. Significant advances have been made in adsorbent materials development and the production cost and uranium capacity of these materials under controlled laboratory settings have brought the theoretical cost of seawater mining within an order of magnitude of terrestrial sourcing. However, a significant unknown variable has been the impact that fouling might have on uranium uptake and recovery and on the reusability of the adsorbents.

An attempt was made to examine where fouling organisms might grow and adhere on the adsorbent fibers and to investigate whether the growth of organisms on the adsorbent resulted in any physical damage to the fibers. Xray microtomography was selected as a means of analysis because it provided a means for both cross-sectional (2-D) and three-dimensional visualization of the fibers and biofilms at appropriate scales and has been previously demonstrated in the analysis of biofilm structures18, 19. For these experiments, the model biofilm and fouling bacterium Pseudomonas fluorescens was grown in LB medium with kanamycin (50 µg/mL) overnight at 37 °C in an incubator shaker. A 9 mL sample of overnight culture was added to a sterile 50 mL centrifuge tube containing the ORNL AF1 adsorbent and incubated for 3 days. The fiber samples were then collected and fixed using 4 % paraformaldehyde (16

The ocean is an extremely harsh environment for the deployment of manmade objects, such as sensors, buoys, filtration systems, remotely operated underwater vehicles (ROVs), and ships. While biofouling and corrosion of solid metallic and painted surfaces have been the subject of many studies, there is comparatively little data currently available for non-metallic, fibrous materials, such as the uranium adsorbents. Marine biofouling occurs as an organic layer of cells, molecules, detritus, and inorganic precipitates that may form a barrier between surfaces and the fluid environment, thus reducing the rate of uranium uptake or the total capacity of the adsorbents. In particular, biomolecular and cellular fouling may invade and block the nano- and micro-scale porosity that is a key feature of high BET surface area materials. These layers may also interfere with recovery processes by diluting or limiting the penetra-



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

tion of extraction solutions. The fouling layer also changes oxygen gradients and underlying fouling organisms can chelate metal ions or greatly influence the surrounding pH and alter redox reaction pathways that may interfere with uranium uptake or lead to accelerated corrosion of the adsorbent material 21, 22. Fouling can be very difficult to remove, with some organic adhesives exceeding the bond strength of commercial cyanoacrylate (i.e., super glue) 23. Because the primary effort of US, Japanese, and Chinese seawater uranium programs has focused on improving the kinetics and total capacity of adsorbent materials, a series of experiments was designed and completed with a focus on testing the impact of biofouling on uranium uptake by adsorbents. The experiments considered conditions that may promote or limit the rate of fouling, developed a correlation between the amount of fouling and the degree of impact, and attempted to place an upper bound on the potential impact fouling may have on the rate of uranium uptake and total capacity of a model adsorbent material. A leading adsorbent material provided by ORNL, AF1 adsorbent, which is based upon a polyethylene trunk material and amidoxime ligands was used for the tests. Current plans call for placing braids of adsorbent material in water < 50 m deep where temperatures are typically warmer and because the potential deployment and recovery costs increase with depth1. However, in many locations this depth is within the photic (euphotic) zone and the rate and extent of biofouling in the presence of light is often greater than that found in the disphotic (twilight) or aphotic zones.24-27 We first analyzed the effect of preestablished fouling on adsorbent fibers with light present (a worst case scenario) under tightly controlled seawater flow conditions that allowed for highly accurate measurements of uranium uptake relative to the amount of passing seawater. A subsequent set of experiments then compared effects on adsorbents with no prior fouling. The tests with and without light simulated how deployment in or below the photic zone might impact uranium uptake. The first round of tests compared adsorbent fibers that were either pre-fouled with N. incerta or untreated prior to packaging in columns. Pre-fouling was used in an attempt to maximize the amount of fouling on the fiber and therefore assess the upper bound of potential impact. The untreated fibers were kept dark while the pre-fouled fiber columns were exposed to light during the seawater exposure period. The 42-day exposure extended from late spring to early summer, May 13 to June 30, 2014. Light was used to simulate shallow water (< 50 m) environments where photosynthetic organisms prolifereate. Photosynthetic species of bacteria and eukaryotes, such as diatoms and macroalgal species (e.g, Ulva spp.) are important contributors to the early stages of biofouling in illuminated settings. 10, 25-28

The seawater used in these exposures was filtered to 0.45 µm to prevent clogging of the columns and disruption of the highly controlled flow regimes. However, we observed the presence of some diatoms (>2 µm) on surfac-

Page 6 of 13

es exposed to the post-filtered water, indicating that some larger eukaryotic species passed through the filtration in small numbers in addition to the smaller prokaryotic species we expected to pass through more easily. Pre-fouling and exposure to light reduced uranium uptake by the AF1 adsorbent 20% after 42 days of adsorption in seawater compared to the untreated, aphotic exposure (Fig. 5). Light alone was not found to affect adsorption in prior controlled experiments using artificial seawater in closed systems with no cells added (personal communication with ORNL adsorbent developers).

Figure 5. Uranium adsorption results from column experiment 1. Filled circles show measured values at different timepoints. The amount of uranium adsorbed by pre-fouled, illuminated fiber (blue circle) was compared to that of untreated, aphotic fiber (red circle). The solid line shows the amount of adsorption calculated by one-site ligand saturation modeling.

To validate the uranium uptake results from the first experiment, a second set of experiments with AF1 adsorbent was performed the following winter using similarly prepared columns, pre-treatment, and the presence of light. Due to materials and sampling limitations caused by other experiments, only 3 samples were collected after 0, 21, and 42 days of seawater exposure from January 29 through March 12, 2015. In the second experiment, uranium adsorption was also decreased 20%, confirming the results of the first test (Fig. 6). In both experiments, visual observation and other analyses found considerably more biological growth associated with the pre-fouled and lightexposed fibers than on the untreated, aphotic fibers (discussed below). To assess the impact light had on biological growth on the samples and whether or not the pre-fouling remained on the samples following the 42-day adsorption period, a number of assays were performed to visualize, characterize, and quantify any fouling material. The most obvious difference between the samples in the first two experiments was the presence of dark coloration on the light-exposed samples (Fig. 7). The columns and fibers were exposed to 0.45 µm filtered seawater, which is greatly reduced in cell content, but not sterile. Smaller marine mi-


Page 7 of 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


croorganisms would have been able to pass through the filter and into the columns where the continuous access to nutrients (e.g., dissolved organic matter) and light would have allowed them to grow. Indeed some color change is visible on the dark enclosed fibers (top panel, Fig 7), but much less than on the light-exposed fibers. This brown coloration is frequently observed on surfaces in our seawater systems in association with the growth of microalgae species.

Figure 6. Uranium adsorption results from column experiment 2. Amount of uranium adsorbed by ORNL AF1 adsorbent after being exposed to filtered seawater for 42 days in the presence of light. Fouling samples were collected at 3 different time points (0, 21, and 42 days; blue line) and compared to non-fouled AF1 adsorbent (red line, the same data shown in Fig. 5).

Figure 7. Visible evidence of enhanced biofouling by light. Column pictures at different time points showed that light enhanced a color change we attribute to the increase in biofouling directly observed by microscopy, TOC, and DNA evidence.

To confirm that the observed color change was associated with enhanced biological growth, plain light microscopy was used to detect the presence of pigmented cells. (Fig. 8). Light microscopy is one of the oldest methods used in the analysis of fouling and biofilms, and remains frequently used as a means to observe the presence or absence of cells or for characterization and enumeration 29, 30 . The image analysis and identifications confirmed that 1) the addition of light resulted in the presence of more diatoms and other microalgae, and 2) that the observed

growth was from native species found in the Sequim Bay seawater and not from pre-fouling with N. incerta. The conditions were so favorable for the native organisms that they outcompeted the introduced diatom species. Not unexpectedly, the results from column experiments demonstrated that the addition of light promoted the growth of eukaryotic phototrophic species. Furthermore, these results suggest that the amount or diversity of biofouling species on adsorbents in the open ocean could be even greater where there is no pre-filtration.

Figure 8. Microscopic evidence of enhanced biofouling by light. Microscopic pictures (1000 X magnification) also showed enhanced biofouling by algal species in the presence of light. A variety of algae (circled) with different morphologies were found on the fibers. A scale bar is not provided due to a technical malfunction.

To further characterize and measure the potential increase in biofouling found on the adsorbent, nonpurgeable organic carbon (NPOC) analysis was performed to quantify the build-up of organic derived carbon on the AF1 adsorbent in the presence or absence of light. NPOC analysis is well-established (e.g., by EPA SW-846 Method 9060)31 for a variety of samples, including water, soil, sediments, and other matrices in which biological material may be found.32-35 We have developed a method for NPOC analysis of biofilms and biofouling on solid, porous, or fibrous matrices that provides for increased extraction efficiency over standard NaOH-based methods while minimizing mineralization (acidification of organic compounds to CO2). The method, which is the subject of a separate publication, has been validated for precision and reproducibility using known quantities of organic carbon standards and outperforms commonly used solvents and extraction protocols. Using standard methods,31-35 the primary issue is underrepresentation of how much biomass is present due to inefficient extraction. However, this has little bearing on making relative comparisons between samples so long as the use of a rigorously standardized protocol provides consistent error rates and high reproducibility. Samples were exposed during the second column experiment. Three columns, each containing 50 mg of untreated fibers (no pre-fouling) were kept in darkness during the 0, 21, and 42-day seawater exposure. A second set of 3 columns, each containing 50 mg each of untreated fibers



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

were exposed to light during the same seawater exposure period. Due to the scarcity of the adsorbent fiber samples and the need to conduct other fouling and chemical analyses, we were limited to a single fiber sample for each time point. The results for each timepoint were generated by consuming the entire 50 mg sample taken from each column (light or dark).

Page 8 of 13

diverse assortment of organisms, but with time a smaller number of taxa come to dominate the surfaces. Since organisms that attached to fibers depended on environmental factors, a better understanding of biofouling community structure will be valuable to develop anti-fouling strategies.

After extraction of the organic carbon was completed as described in the Experimental Methods section, the Shimadzu TOC-L analyzer generated a dilution series from each sample and analyzed each fraction separately to generate a curve and to calculate the amount of TOC present. Results from this experiment clearly showed that at 21 and 42 days, the amount of organic carbon found on the light exposed fibers is higher than that found on the dark exposed fibers at both time points; equating to a 29 % and 27 % difference at 21 and 42 days, respectively (Fig. 9). NPOC analysis supports the visual observation and microscopy results, confirming that a large portion of the fouled surface was derived from organic carbon, not inorganic carbonates or other minerals associated with seawater.

Figure 9. NPCO analysis of AF1 adsorbent. The amount of biomass accumulated on AF1 adsorbent was increased in the presence of light.

Lastly, DNA analysis of the fouled AF1 adsorbent was completed to characterize the microbial community composition associated with the fouled adsorbent. Cells were harvested from the fibers and total genomic DNA was extracted from the cells. A PCR based approach was used to amplify a portion of the genome, the length of which may be used to distinguish different taxa from each other. In Figure 10, the X-axis equates to the length of the DNA and the Y-axis is a relative measure of quantity. Each peak represents unique fragment length or microbial taxon. This analysis showed that the microbial communities involved in biofouling of the adsorbent fibers are diverse (multiple taxa) and that these communities evolve; the community composition changes with exposure time and illumination (Fig. 10). This supports the microscopy results that observed the presence of a diverse group of diatoms attached to the fibers. The pattern in both the dark and light exposed fibers suggests that the surfaces are initially colonized by a

Figure 10. Impact of light on microbial communities involved in biofouling. Light impacts the diversity of microbial community involved in biofouling. (A) Dark column 21 days (B) Dark col-


Page 9 of 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


umn 42 days (C) Light column 21 days (D) Light column 42 days. Each peak represents a unique DNA sequence and therefore, indicates unique microbial community structure, which varies by exposure to light and time. Peaks are labeled by size in base pairs (BP).

Effect of biofouling on uranium uptake by braided adsorbent materials using flume testing The leading envisioned deployment strategy for uranium adsorbents is to weave the adsorbent fibers into long braids that would be anchored to the seafloor and allowed to undulate in the local ocean current like artificial blades of kelp 2, 36. This format may result in uneven water circulation across fibers that are surface exposed or located within the braid and movement over the braids might allow the fibers to rub against each other to remove fouling material. Following the column studies, an experiment with regular and mini braids was completed using flumes to assess how biofouling might affect adsorbent performance using the ‘kelp bed’ model of deployment. Flume testing allowed the adsorbents to freely move in seawater, while adsorbents were immobilized in column settings. It was unknown how the movement of adsorbent fiber may affect biofouling. Additionally, the flume test was done with 150 µm filtered seawater just to remove large particles. ORNL AI8 adsorbent was conditioned and exposed to seawater for 42 days either in the presence or absence of simulated sunlight. Control adsorbents were placed in columns and exposed to 0.45 µm filtered seawater in the absence of light. Adsorbents in the illuminated flume resulted in up to 30 % less uranium adsorption compared to control samples that were exposed to filtered seawater in dark columns (Fig. 11). Adsorbents in the dark flume, however, showed minimal adsorption loss by biofouling (Fig. 11).

Figure 11. Uranium uptake results from flume experiment. Light flume: Uranium adsorption of ORNL AI8 adsorbent after being exposed to 150 µm filtered seawater for 42 days in the presence of light. Samples were collected at 6 different time points. Dark Flume: AI8 adsorbent was exposed to 150 µm filtered seawater in dark. Samples were collected after 42 days. Control column: AI8L2R2 was exposed to filtered seawater in dark. Samples were collected after 42 days. Reference: Previous time series of

dark column exposure experiment with a previous batch of ORNL AI8 adsorbents.

The impact of light on biofouling was also observed in the flume experimental setup (Fig. 12), and the visible effects of biofouling appeared to be even greater than was seen in the column experiments. Heavy growth was found on the walls of the flume and on the braided fibers. This was likely because the adsorbents were exposed to a greater number and diversity of organisms in the coarsely filtered seawater used in the flumes. Additionally, the structure of the braid provided different textures and crevices between the fibers where microorganisms could colonize.

Figure 12. Impact of light on biofouling. (A) Flume experiment Day 1 (B) Flume experiment Day 42: Considerable amount of biofouling occurred in light flume compared to dark flume.

The adsorbents in the light flume increased in dry mass by approximately 80 % after 42 days of exposure to 150 µm filtered seawater at 20 °C (Fig. 13). The adsorbents in the dark flume, on the other hand, only increased 20 % over the same exposure period, exposed to identical source seawater and temperature (Fig. 13). The braid and flume experiment data suggest that biofouling in sunlit shallow seawater has the potential to reduce uranium adsorption capacity by ~30 % after 42 days of exposure. Minimal or no adsorption loss due to biofouling occurred in the dark flume exposure as evidenced by a similar response between adsorption in the flume and the AI8 adsorption material that underwent 32 day testing in a flow-through columns.

Figure 13. Increase in the dry mass of biofouling material as a function of time in illuminated and dark flumes. The accumulation of biofouling material over a 42-day period is represented as biofouling mass (g) per adsorbent mass (g).

Enhanced biofouling by light was also evident using microscopy (Fig. 14 and 15). Both flume and microscope images visually demonstrate that controlling light



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

exposure can substantially reduce biofouling from photosynthetic organisms.

Figure 14. AI8 adsorbent in dark flume. Braided AI8 adsorbent samples were collected at 6 different time points and examined under a microscope (1000 X magnification). Scale bar represents 100 µm. Very few algal cells were visible on the fibers.

Page 10 of 13

vide an unprecedented 3-D visualization of the fibers and also demonstrated that growth of P. fluorescens did not readily damage the structure of the fibers. This was the first time that X-ray microtomography had been used to examine any of the uranium adsorbents and the images have provided details of the 3-D structure in an aqueous suspension that had been difficult to acquire previously. The inclusion of an aqueous medium is critical as the 3-D structure of many of the adsorbents collapses under dry conditions (unpublished data). Scheduling at the APS limited the amount of preparative time available before the imaging was performed, and although P. fluorescens grows rapidly, the 3-day exposure on the fiber may not have been sufficient to allow for good surface adhesion, colonization, and early stage biofilm formation. Additionally, the preparative steps may have dislodged cells from the surface of the fiber; it was necessary to immobilize the fibers prior to imaging. Visual observation of the fiber did not show obvious signs of fouling, such as discoloration. Therefore, it is inconclusive whether the APS is a suitable technology for imaging early stage microbial fouling on such fibers.

Figure 15. AI8 adsorbent in light flume. AI8L2R2 braid samples were collected at 6 different time points and examined under a microscope (1000 X magnification). Scale bar represents 100 µm. Many algal cells were observed attached to the adsorbent fibers.

Imaging fouled adsorbent fiber To better understand where biofouling occurs on the fibers and what impact it may have on the exposed surface, fouled fibers were imaged using the advanced photon source (APS) at Argonne National Laboratory. ORNL AF1 adsorbent fibers were exposed to Pseudomonas fluorescens, a gram-negative bacterium used as a model organism to study the formation of biofilms. The goal was to induce biofilm formation on the surface of the fiber that could be resolved from the structural components of the fiber using the imaging capability of the APS Beamline 2BM. A control fiber sample that was only exposed to sterile culture medium was also imaged. Figure 16 shows a 3D rendered image of two P. fluorescens exposed fibers that illustrates their structure. The color is related to the x-ray attenuation and depends strongly on average atomic number and a bit less strongly on density. Coloration was performed on a continuous scale, therefore individual features were not identified and then colored. Despite the dramatic images of the fiber structures, biofilms and cells were not clearly observed in the images and no differences were seen between the fouled and control sample. The images did, however, pro-

Figure 16. 3D image of ORNL AF1 adsorbent. Fiber structure is illustrated. The ellipsoidal cross-section of the fiber is approximately 100 µm by 200 ~ 300 µm.

CONCLUSION PNNL has tested the potential impact of biofouling on adsorption, capacity, and kinetics of amidoximebased uranium adsorbent materials. Biofouling includes the unwanted accumulation of both cellular and biomolecular components (conditioning film and cell-generated) onto the surface of the adsorbents. Results from these biofouling experiments illustrate the importance of reducing exposure to light and indicate that fouling formation on the adsorbent reduces the rate of uranium adsorption and total adsorbent capacity. Assessment of fiber performance was determined using ORNL AF1 and AI8 adsorbents in loose fiber or braided formats and in column or flume settings, respectively. Tests with all fibers and test beds were performed in the presence or absence of light to simulate shallow (photic zone) and deep seawater (aphotic zone) deployment conditions. Testing these fibers under sunlit shallow seawater


Page 11 of 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


conditions indicates that the accumulation of biomass (e.g., through biological growth) interfered with the adsorption of uranium on the fibers. As the amount of biofouling increased on the fiber, a corresponding decrease in the ability of the fiber to uptake uranium was also observed. Even with low cell density filtered seawater, the adsorbent materials were highly susceptible to cellular colonization and were negatively affected by biofouling. Therefore, the effects of biofouling must be considered in the future design, construction, and deployment strategy of adsorbent materials. Ideally it would have been possible to set up experiments in which adsorbents were exposed to sterile seawater as a perfect negative control, but it was not possible to generate the large volume of sterile seawater necessary. UV, chemical, or autoclave sterilization would all introduce unique variables into the seawater and ultrafiltration was not feasible at the necessary scale. However, the comparative studies between light and dark exposed adsorbents and the negative impact of light on uranium uptake demonstrate the importance of deploying the adsorbent below the photic zone. Furthermore, the multiple lines of evidence from the visual observations, microscopy, and NPOC analysis all indicate that exposure to light increased the amount of fouling on the surface of the fibers and this is likely the cause of the interference. Development of next-generation adsorbents that exhibit antifouling and/or self-cleaning (grooming) properties would help to mitigate the effects of fouling. As adsorbent fibers are made to create more surface area to promote uranium adsorption, they also have the unwanted effect of enhancing surface contact with fouling organisms and biomolecules. Furthermore, it is anticipated that effects of completely unfiltered seawater will only further promote biofouling of the adsorbent materials, potentially surpassing the 30 % uranium reduction exhibited in the flume experiments (150 µm filtration). The development and testing of future adsorbent fibers should not only consider theoretical adsorption limits determined in a laboratory setting, but also strongly consider how environmental factors will influence adsorbent performance when placed in a marine system. The results of these experiments demonstrate that fouling can significantly reduce uranium uptake. In a high fouling setting, a strategy may need to be employed to prevent fouling, or deployments will need to weigh the benefit of prolonged exposures intended to increase uranium uptake with the corresponding buildup of fouling material and its negative impact. The studies also demonstrate that locating adsorbents below the photic zone may greatly reduce fouling, but deployment in deeper water may add operational costs and deeper waters are also typically cooler, which would interfere with the adsorption kinetics of most adsorbents that favor warmer temperatures. Fouling effects on uranium uptake and the cost of mitigation strategies must be factored into any economic analysis of a seawater uranium program. Analysis of the effect of fouling on uranium ex-

traction processes and on the durability and reuse of fibers will further refine the economic models and conceptual system of operation. Fortunately, many strategies exist to combat fouling and to limit its effects; empirical testing is necessary to provide the data needed to make a cost benefit analysis and to determine the best combination of deployment location, exposure time, and mitigation. AUTHOR INFORMATION Corresponding Author Phone: 360-681-3678 Email: [email protected] Phone: 360-582-2528; Fax: 360-681-4559; E-mail: [email protected] ACKNOWLEDGMENTS This work was supported by the U.S. Department of Energy under Contract DE-AC05-76RL01830 and the Pacific Northwest National Laboratory’s Chemical Imaging Initiative (CII). This research also used resources of the Advanced Photon Source, a U.S. Department of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by Argonne National Laboratory under Contract No. DE-AC02-06CH11357. The authors would like to thank Dana L. Woodruff for identification of organisms from 150 µm filtered seawater and Matthew J. Marshall for acquiring APS imaging data. We also wish to acknowledge the contributions of Yatsandra Oyola, Sadananda Das, and Richard T. Mayes, all of ORNL, in the production of the adsorbent fibers used for the tests. ABBREVIATIONS MSL; Marine Science Laboratory, NPOC; Non-Purgeable Organic Carbon, ORNL: Oak Ridge National Laboratory, PCR; polymerase chain reaction, PNNL; Pacific Northwest National Laboratory REFERENCES 1. DOE Nuclear Energy Research and Development Roadmap: Report to Congress; 2010. 2. Kim, J.; Tsouris, C.; Oyola, Y.; Janke, C. J.; Mayes, R. T.; Dai, S.; Gill, G.; Kuo, L.-J.; Wood, J.; Choe, K.-Y.; Schneider, E.; Lindner, H., Uptake of Uranium from Seawater by Amidoxime-Based Polymeric Adsorbent: Field Experiments, Modeling, and Updated Economic Assessment. Industrial & Engineering Chemistry Research 2014, 53, (14), 6076-6083. 3. Hills, J. M.; Thomason, J. C., The effect of scales of surface roughness on the settlement of barnacle (Semibalanus balanoides) cyprids. Biofouling 1998, 12, (13), 57-69. 4. Berntsson, K. M.; Jonsson, P. R.; Lejhall, M.; Gatenholm, P., Analysis of behavioural rejection of microtextured surfaces and implications for recruitment by the



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

barnacle Balanus improvisus. Journal of Experimental Marine Biology and Ecology 2000, 251, (1), 59-83. 5. Callow, M. E.; Jennings, A. R.; Brennan, A. B.; Seegert, C. E.; Gibson, A.; Wilson, L.; Feinberg, A.; Baney, R.; Callow, J. A., Microtopographic Cues for Settlement of Zoospores of the Green Fouling Alga Enteromorpha. Biofouling 2002, 18, (3), 229-236. 6. Howell, D.; Behrends, B., A methodology for evaluating biocide release rate, surface roughness and leach layer formation in a TBT-free, self-polishing antifouling coating. Biofouling 2006, 22, (5), 303-315. 7. Abarzua, S.; Jakubowski, S.; Eckert, S.; Fuchs, P., Biotechnological Investigation for the Prevention of Marine Biofouling II. Blue-Green Algae as Potential Producers of Biogenic Agents for the Growth Inhibition of Microfouling Organisms. In Botanica Marina, 1999; Vol. 42, p 459. 8. Lejars, M.; Margaillan, A.; Bressy, C., Fouling Release Coatings: A Nontoxic Alternative to Biocidal Antifouling Coatings. Chemical Reviews 2012, 112, (8), 4347-4390. 9. Callow, M. E.; Fletcher, R. L., The influence of low surface energy materials on bioadhesion — a review. International Biodeterioration & Biodegradation 1994, 34, (3–4), 333-348. 10. Walt, D. R.; Smulow, J. B.; Turesky, S. S.; Hill, R. G., The effect of gravity on initial microbial adhesion. Journal of Colloid and Interface Science 1985, 107, (2), 334-336. 11. Chambers, L. D.; Stokes, K. R.; Walsh, F. C.; Wood, R. J. K., Modern approaches to marine antifouling coatings. Surface and Coatings Technology 2006, 201, (6), 3642-3652. 12. de Messano, L. V. R.; Sathler, L.; Reznik, L. Y.; Coutinho, R., The effect of biofouling on localized corrosion of the stainless steels N08904 and UNS S32760. International Biodeterioration & Biodegradation 2009, 63, (5), 607-614. 13. Champ, M. A., A review of organotin regulatory strategies, pending actions, related costs and benefits. Science of The Total Environment 2000, 258, (1–2), 21-71. 14. Yebra, D. M.; Kiil, S.; Dam-Johansen, K., Antifouling technology—past, present and future steps towards efficient and environmentally friendly antifouling coatings. Progress in Organic Coatings 2004, 50, (2), 75104. 15. Abbott, A.; Abel, P. D.; Arnold, D. W.; Milne, A., Cost–benefit analysis of the use of TBT: the case for a treatment approach. Science of The Total Environment 2000, 258, (1–2), 5-19. 16. Cooney, J. J.; Tang, R.-J., [47] Quantifying effects of antifouling paints on microbial biofilm formation. In Methods in Enzymology, Academic Press: 1999; Vol. Volume 310, pp 637-644. 17. Cao, S.; Wang, J.; Chen, H.; Chen, D., Progress of marine biofouling and antifouling technologies. Chinese Science Bulletin 2011, 56, (7), 598-612. 18. Davit, Y.; Iltis, G.; Debenest, G.; Veran-Tissoires, S.; Wildenschild, D.; Gerino, M.; Quintard, M., Imaging biofilm in porous media using X-ray computed

Page 12 of 13

microtomography. Journal of Microscopy 2011, 242, (1), 15-25. 19. Erin A. Miller, X. X., Micah Miller, Paul Keller, Timothy A. White, Matthew Marshall, Investigating biofilm structure using X-ray microtomography and gratings-based phase contrast. DEVELOPMENTS IN XRAY TOMOGRAPHY VIIL 2012, 8506. 20. De Carlo, F.; Xiao, X.; Tieman, B. In X-ray tomography system, automation, and remote access at beamline 2-BM of the Advanced Photon Source, 2006; 2006; pp 63180K-63180K-13. 21. Kinzler, K.; Gehrke, T.; Telegdi, J.; Sand, W., Bioleaching—a result of interfacial processes caused by extracellular polymeric substances (EPS). Hydrometallurgy 2003, 71, (1–2), 83-88. 22. Beech, I. B.; Sunner, J., Biocorrosion: towards understanding interactions between biofilms and metals. Current Opinion in Biotechnology 2004, 15, (3), 181-186. 23. Tsang, P. H.; Li, G.; Brun, Y. V.; Freund, L. B.; Tang, J. X., Adhesion of single bacterial cells in the micronewton range. Proceedings of the National Academy of Sciences of the United States of America 2006, 103, (15), 5764-5768. 24. Rao, T. S.; Rani, P. G.; Venugopalan, V. P.; Nair, K. V. K., Biofilm formation in a freshwater environment under photic and aphotic conditions. Biofouling 1997, 11, (4), 265-282. 25. Onuf, C., Biofouling and the continuous monitoring of underwater light from a seagrass perspective. Estuaries and Coasts 2006, 29, (3), 511-518. 26. Zecher, K.; Jagmann, N.; Seemann, P.; Philipp, B., An efficient screening method for the isolation of heterotrophic bacteria influencing growth of diatoms under photoautotrophic conditions. Journal of Microbiological Methods 2015, 119, 154-162. 27. Cao, S.; Wang, J.; Chen, D., Influence of Illumination on Settlement of Diatom Navicula sp. Microbial Ecology 2011, 62, (4), 931-940. 28. Cooksey, K. E., Wigglesworth-Cooksey, B., Adhesion of bacteria and diatoms to surfaces in the sea; a review. Aquatic Microbial Ecology 1995, 9, 87-96. 29. Walker, J. T., Keevil, C.W., Study of micobial biofilms using light microscope techniques. International Biodeterioration & Biodegradation 1994, 34, (3-4), 223~236. 30. Wolf, G.; Crespo, J.; Reis, M. M., Optical and spectroscopic methods for biofilm examination and monitoring. Reviews in Environmental Science and Biotechnology 2002, 1, (3), 227-251. 31. EPA, Total organic carbon (TOC) in soil. In Agency, E. P., Ed. 1999. 32. Favilli, F.; Egli, M.; Cherubini, P.; Sartori, G.; Haeberli, W.; Delbos, E., Comparison of different methods of obtaining a resilient organic matter fraction in Alpine soils. Geoderma 2008, 145, (3–4), 355-369. 33. Mikutta, R.; Kleber, M.; Kaiser, K.; Jahn, R., Review. Soil Science Society of America Journal 2005, 69, (1).


Page 13 of 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


34. Clark, K. A. Total organic carbon analysis for cleaning validation in pharmaceutical manufacturing; 2000. 35. Schumacher, B. A., Methods for the determination of total organic carbon (TOC) in soils and sediments. Ecological Risk Assessment Support Center 2002, 2002, 123. 36. Tamada, M. In Current Status of Technology for Collection of Uranium from Seawater, Erice International Seminars on Planetary Emergencies, Erice, Italy, 2009; Erice, Italy, 2009.

Insert Table of Contents artwork here


Effect of Biofouling on the Performance of ... - ACS Publications

Recommend Documents