Extracellular polymeric substances govern the surface charge of biogenic elemental selenium nanoparticles.

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Extracellular polymeric substances govern the surface charge of biogenic elemental selenium nanoparticles Rohan Jain Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/es5043063 • Publication Date (Web): 23 Dec 2014 Downloaded from http://pubs.acs.org on December 27, 2014

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Extracellular polymeric substances govern the surface charge of biogenic elemental selenium nanoparticles

Journal: Manuscript ID: Manuscript Type: Date Submitted by the Author: Complete List of Authors:

Environmental Science & Technology es-2014-043063.R1 Article 22-Dec-2014 Jain, Rohan; UNESCO-IHE, EEWT Jordan, Norbert; Helmholtz-Zentrum Dresden-Rossendorf Institute of Resource Ecology, Weiss, Stephan; Helmholtz-Zentrum Dresden-Rossendorf, Institute of Resource Ecology Foerstendorf, Harald; Helmholtz-Zentrum Dresden-Rossendorf e.V., Institute of Resource Ecology Heim, Karsten; Helmholtz-Zentrum Dresden-Rossendorf e.V., Institute of Resource Ecology Kacker, Rohit; Delft University of Technology, Process & Energy Laboratory Huebner, Rene; Helmholtz-Zentrum Dresden-Rossendorf e.V., Institute of Ion Beam Physics and Materials Research Kramer, Herman J.M.; Delft University of Technology, Process & Energy Laboratory van Hullebusch, Eric; Université Paris-Est, Laboratoire Géomatériaux et Environnement Farges, Francois; Muséum National d’Histoire Naturelle, Laboratoire de Minéralogie et de Cosmochimie du Muséum Lens, Piet N; UNESCO-IHE Institute for Water Education, Environmental Resources Department

ACS Paragon Plus Environment

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Manuscript submitted to Environmental Science & Technology

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Extracellular polymeric substances govern the surface charge of biogenic elemental selenium nanoparticles

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Rohan Jain1,5*, Norbert Jordan2, Stephan Weiss2, Harald Foerstendorf2, Karsten Heim2, Rohit Kacker3, René Hübner4, Herman Kramer3, Eric D. van Hullebusch5, François Farges6, Piet N. L. Lens1

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UNESCO-IHE, Institute for Water Education, Westvest 7, 2611AX Delft, The

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Netherlands

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2

Institute of Resource Ecology, Helmholtz-Zentrum Dresden-Rossendorf e.V.,

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Bautzner Landstr. 400, D-01328 Dresden, Germany

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Process & Energy Laboratory, Delft University of Technology, Leeghwaterstraat 44, 2628 CA Delft The Netherlands

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Institute of Ion Beam Physics and Materials Research

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Helmholtz-Zentrum Dresden-Rossendorf e.V., Bautzner Landstr. 400, D-01328

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Dresden, Germany

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Université Paris-Est, Laboratoire Géomatériaux et Environnement (EA 4508), UPEM, 77454 Marne la Vallée, France

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Institut de Mineralogie, de Physique des Materiaux et de Cosmochimie (IMPMC),

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Muséum National d'Histoire Naturelle, Université Pierre-et-Marie Curie and CNRS UMR

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7590, Paris, France.

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*Corresponding author:


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Manuscript submitted to Environmental Science & Technology 23

Phone: +31 152151816, fax: +31 152122921 e-mail: [email protected], mailto:

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UNESCO-IHE, Institute for Water Education, Westvest 7, 2611AX Delft, The

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Netherlands

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ABSTRACT

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The origin of the organic layer covering colloidal biogenic elemental selenium

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nanoparticles (BioSeNPs) is not known, particularly in the case when they are

49

synthesized by complex microbial communities. This study investigated the presence of

50

extracellular polymeric substances (EPS) on BioSeNPs. The role of EPS in capping the

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extracellularly avialble BioSeNPs was also examined. FT-IR spectroscopy and

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colorimetric measurements confirmed the presence of functional groups characteristic

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of proteins and carbohydrates on the BioSeNPs, suggesting the presence of EPS.

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Chemical synthesis of elemental selenium nanoparticles in the presence of EPS,

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extracted from selenite fed anaerobic granular sludge, yielded stable colloidal spherical

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selenium nanoparticles. Furthermore, extracted EPS, BioSeNPs and chemically

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synthesized EPS capped selenium nanoparticles had similar surface properties, as

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shown by ζ-potential versus pH profiles and iso-electric point measurements. This study

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shows that the EPS of anaerobic granular sludge forms the organic layer present on the

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BioSeNPs synthesized by these granules. The EPS also govern the surface charge of

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these BioSeNPs, thereby contributing to their colloidal properties, hence affecting their

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fate in the environment and the efficiency of bioremediation technologies.

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.

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Keywords: EPS, BioSeNPs, surface charge, capping, FT-IR

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Table of contents

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INTRODUCTION

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Selenium is an essential nutrient in the human diet.1 However, the higher

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concentrations of selenium, especially those of the selenium oxyanions selenate and

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selenite, are toxic to humans, animals and aquatic life.2–4 Therefore, regulatory

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agencies have set limits on total selenium discharges, e.g. the Environmental Protection

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Agency of the United States has recommended a discharge limit of 5 µg L−1 total

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selenium in freshwater.5 Anaerobic bioreduction of dissolved selenium oxyanions to

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elemental selenium is considered a promising technology for the remediation of

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selenium oxyanions containing wastewaters.6,7 However, the produced biogenic

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elemental selenium is in the form of colloidal spherical nanoparticles with a diameter of

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50 - 500 nm.8,9 Such colloidal biogenic elemental selenium nanoparticles (BioSeNPs) is

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present in high concentrations in the effluent of upflow anaerobic sludge blanket

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reactors (UASB), in which anaerobic granules treat selenium rich wastewaters.7 Buchs

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et al. (2013)10 showed that the colloidal properties of these BioSeNPs determine their

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transport and fate in the environment as well as the bioremediation efficiency. Thus, it is

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important to understand the factors governing the colloidal properties of BioSeNPs.

102 103

Capping agents are known to affect the surface properties of chemically produced

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metal(loid) nanoparticles, including surface charge and consequently colloidal stability

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(Figure S1 in SI).11 For instance, sterically stabilized silver nanoparticles capped by

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polyvinylpyrrolidone (PVP) do not agglomerate while electrostatically stabilized silver

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nanoparticles by citrate do agglomerate at low pH or high ionic strength.12

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Electrosterically stabilized silver nanoparticles by branched polyethyleneimine capping


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are more resistant to agglomeration at low pH or high ionic strength as compared to

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citrate stabilized silver nanoparticles.12 Proteins such as bovine serum albumin (BSA),

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which stabilize silica nanoparticles through electrosteric mechanisms13, are also known

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to stabilize chemically produced selenium nanoparticles (CheSeNPs).14 Proteins are

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also known to be associated with the BioSeNPs.15,16 It has been proposed that

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BioSeNPs are coated with an organic layer of microbial origin, composed not

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exclusively of proteins.17 However, to the best of our knowledge, the origin of this

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organic layer and its effect on the surface charge and thus, on the colloidal properties of

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BioSeNPs, is not known.

118 119

This study hypothesized that extracellular polymeric substances (EPS) are the capping

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agents and thus can affect the surface charge of the BioSeNPs that are available

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extracellularly. EPS are high molecular weight macromolecules that contain mainly

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proteins, carbohydrates, humic-like substances and small concentrations of DNA.18,19

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Thus, they provide many sites that can interact with the elemental selenium. EPS are an

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important component of mixed microbial aggregates i.e. biofilms or anaerobic granules

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employed for the treatment of selenium rich wastewaters in bioreactors.19,20,21 Besides,

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reduction of selenite to BioSeNPs by pure cultures has been reported both in the

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periplasmic space22 and extracellularly23, which further indicates that the BioSeNPs are

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likely to be grown in the presence of EPS.

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In this study, the presence of an organic layer on the surface of BioSeNPs was

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determined from Energy Disperse X-ray spectroscopy (EDXS) and ζ-potential


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measurements as well as acid-base titrations. The presence of proteins and

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carbohydrates, suggesting the presence of EPS on the BioSeNPs, was confirmed by

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Fourier-transform Infrared Spectroscopy (FT-IR) and colorimetric measurements. EPS

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extracted from selenite fed anaerobic granules was used as a capping agent for

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CheSeNPs (EPS capped CheSeNPs) and their colloidal properties were studied as

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well. BSA, a well-known capping agent for CheSeNPs14,24, was used as a reference

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material to (a) demonstrate the capping ability of EPS and (b) show that the capping of

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BioSeNPs does not exclusively consists of proteins.

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MATERIALS AND METHODS

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BioSeNPs production and purification

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BioSeNPs were produced using an anaerobic granular sludge treating pulp and paper

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wastewater, which has been described in detail by Roest et al. (2005)25. Anaerobic

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granular sludge (13 g L−1 wet weight) was added to the oxygen-free growth medium

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(NH4Cl 5.6, CaCl2.2H2O 0.1, KH2PO4 1.8, Na2HPO4 2.0, KCl 3.3, in mM) with 20.0 mM

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of sodium lactate and 5.0 mM of sodium selenite. The incubation was carried out at 30

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°C and pH 7.3 for 14 days. The production of elemental selenium was confirmed by the

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appearance of a red color (Figure S2b in SI). The produced BioSeNPs were purified

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following the protocol developed in Dobias et al.16 with minor modifications. Briefly, the

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supernatant was decanted, followed by simple centrifugation (Hermle Z36HK) at 3,000

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g and 4 °C for 15 minutes to separate the suspended biomass. The collected BioSeNPs

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present in the supernatant from the previous centrifugation step were concentrated by

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centrifugation (Hermle Z36HK) at 37,000 g and 4 °C for 15 minutes. The pellet was re-


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suspended in Milli-Q (18MΩ cm) water and purified by sonication (15 minutes at

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23 KHz, Soniprep 150, UK) followed by hexane separation. The concentration of

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BioSeNPs was determined by dissolving them in concentrated HNO3 and then

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measuring the Se concentration by ICP-MS.26

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Analysis of the chemical composition of the BioSeNPs' surface

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30 mL of purified BioSeNPs (390 mg L−1) were sonicated for 15 minutes at 23 KHz

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using Soniprep 150 (MSE, UK) sonicator. After the sonication, BioSeNPs were

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centrifuged at 37,000 g for 30 minutes at 4 oC (Hermle Z36HK). The supernatant was

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collected and analyzed for carbohydrates (phenol-sulfuric acid method)27, proteins and

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humic-like substances (modified lowry method by Fr¢lund et al., 199528). The DNA

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concentration in the supernatant was measured after precipitation of the DNA with iso-

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propanol and then measuring the absorbance of the pellet using a spectrophotometer at

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260 nm.29

169 170

EPS extraction and characterization

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EPS was extracted from anaerobic granules, which were fed with selenite and lactate,

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and incubated at 30 °C for 14 days, using the NaOH extraction method.30 It is important

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to note that the NaOH extraction method may slightly alter the molecular structure of the

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EPS19, which is, however, unavoidable.

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The total organic carbon (TOC) and total organic nitrogen (TN) content of the extracted

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EPS was determined using a total organic carbon analyzer (Shimadzu TOC-VCPN

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analyzer, Kyoto, Japan). 3D excitation (220 - 400 nm) and emission (300 - 500 nm)


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fluorescent spectroscopy of extracted EPS (total organic carbon concentration 0.5 mg

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L−1) was carried out using a FluoroMax-3 spectrofluorometer (HORIBA Jobin Yvon,

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Edison, NJ, USA) instrument. The carbohydrate, protein, humic-like substances and

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DNA content in the EPS were determined as described above. The FT-IR spectra of

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EPS were recorded on a Bruker Vertex 70/v spectrometer equipped with a D-LaTGS-

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detector (L-alanine doped triglycine sulfate) (more details in SI).

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Production and purification of CheSeNPs, EPS capped CheSeNPs and BSA

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capped CheSeNPs

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CheSeNPs were produced by reduction of sodium selenite (100 mM, 0.35 mL) by L-

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reduced glutathione (GSH) (100 mM, 1.4 mL) in a total volume of 30 mL at 22 °C. EPS

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capped CheSeNPs and BSA capped CheSeNPs (chemically produced selenium

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nanoparticles in the presence of EPS and BSA, respectively) were produced in a similar

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manner, but in the presence of 100 mg L−1 total organic carbon of the extracted EPS

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and 100 mg L−1 of BSA, respectively. After the addition of NaOH (1 M) to adjust the pH

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to 7.2, the produced CheSeNPs, EPS capped CheSeNPs and BSA capped CheSeNPs

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were dialyzed against Milli-Q water (18MΩ cm) using a 3.5 kDa regenerated cellulose

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membrane while changing water every 12 hours for 96 hours.14

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Selenium nanoparticles characterization

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BioSeNPs were characterized by scanning electron microscopy (SEM) coupled with

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EDXS, ζ-potential measurements, hydrodynamic diameter (HDD) measurements, FT-IR

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and acid-base titrations (see SI for more details). CheSeNPs were characterized by


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transmission electron microscopy (TEM) coupled with EDXS (TEM-EDXS) (see SI for

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more details). BSA and EPS coated CheSeNPs were characterized by TEM-EDXS, ζ-

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potential measurements, HDD measurements and FT-IR spectra (see SI for more

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details). EPS and BSA capped CheSeNPs were contacted with different initial

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concentrations of Zn for ζ-potential measurements (see SI for more details).

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RESULTS

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SEM-EDXS analysis of BioSeNPs

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The red BioSeNPs synthesized by the reduction of SeO32− by anaerobic granular sludge

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are primarily spherical in shape (Figure S2a in SI). The resultant BioSeNPs formed a

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stable colloidal suspension (Figure S2b in SI). EDXS analysis of the BioSeNPs

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confirmed the presence of selenium (Figure S2c in SI). In addition, carbon, nitrogen,

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oxygen as well as weak signals of phosphorous, sulfur, calcium, and iron were also

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observed. The presence of carbon, nitrogen, oxygen, phosphorous and sulfur may be

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attributed to the EPS coating of the BioSeNPs, while calcium and iron can be likely

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traced back to the anaerobic granules used for the production of BioSeNPs.

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Determination of functional groups present on the surface of BioSeNPs

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The carbohydrates, proteins, humic-like substances and DNA concentrations released

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in the supernatant after the sonication of purified BioSeNPs were, respectively, 313.8 ±

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3.5, 144.1 ± 2.1, 158.2 ± 2.3 and 4.6 ± 0.8 mg g−1 of BioSeNPs, confirming the

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presence of EPS components19 on the surface of the BioSeNPs.


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Acid-base titrations were carried out to determine the pKa values of the different

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functional groups present on the surface of the BioSeNPs. The acid-base titration

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curves for BioSeNPs (Figure S3 in SI) showed smoother lowering of the pH, as

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compared to the control, with the addition of acid (HCl). This can be attributed to the

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buffering capacity of the BioSeNPs due to the presence of various functional groups on

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their surface.31

229 230

To evaluate the buffering capacity of BioSeNPs in more detail, the derivative of the acid-

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base titration was plotted with pH (Figure 1). The local minima represent the minimum

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variation of the pH and hence the buffering zones due to the adsorption of H+ ions on

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the surface of BioSeNPs, which corresponds to pKa values of the functional groups

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present on the surface of the BioSeNPs.32 It is important to note that local minima will

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not be observed prominently in the beginning and the end of titration due to the

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relatively small pH change. The different pKa values and their corresponding functional

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groups are detailed in Figure 1. The presence of carboxylic acid (pKa = 3.9), phosphoric

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groups (pKa = 6.3) and sulfonic, sulfinic or thiol groups (pKa = 7.5) were confirmed by

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the acid-base titration (Figure 1).31 Most notable absents are the amino groups, but this

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may be due to a small change in the pH value at the beginning of the titrations.

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Figure 1. Derivative of acid-base titration data of BioSeNPs (—).

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Characterization of EPS, EPS capped CheSeNPs, BSA capped CheSeNPs and

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CheSeNPs

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The total organic carbon and total nitrogen concentration of the extracted EPS was

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116.7 ± 0.5 mg L−1 and 19.0 ± 0.3 mg L−1, respectively. 3D excitation emission

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fluorescent spectroscopy of EPS confirmed the presence of aromatic proteins and

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humic substances (Figure S4 in SI). The carbohydrates, proteins, humic-like substances

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and DNA concentrations in the extracted EPS were, respectively, 106.9 ± 2.3, 239.5 ±

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6.2, 184.7 ± 15.3 and 2.7 ± 0.6 mg L−1.

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The extracted EPS and BSA were used as a capping agent for CheSeNPs produced by

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the reduction of sodium selenite using reduced-glutathione.14 The addition of EPS

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during the chemical synthesis of selenium nanoparticles led to the formation of a clear

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suspended yellow-red colloidal solution (Figure 2a). A comparable result was obtained

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for BSA stabilized CheSeNPs (Figure 2b). In contrast, selenium nanoparticles produced

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in the absence of EPS formed a turbid brownish suspension (Figure 2c) in an hour, but

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settled only within two days. No agglomeration and settling of EPS and BSA capped


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CheSeNPs were observed even two weeks after their formation. It is important to note

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that the selenium concentration in the BSA capped CheSeNPs, EPS capped

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CheSeNPs and CheSeNPs was identical (1.1 mM). TEM micrographs revealed that

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EPS and BSA capped CheSeNPs were spherical in shape (Figure 2d, e). In contrast,

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wires of selenium were formed in the absence of EPS (Figure 2f). Obviously, EPS acted

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as a capping agent, thereby influencing the morphology of the CheSeNPs.33–35 EDX

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spectra confirmed the presence of selenium in all three types of CheSeNPs investigated

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(Figure 2g, h, i).

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Figure 2. CheSeNPs formed in the presence of the capping agents (a) EPS and (b)

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BSA as well as (c) in the absence of any capping agent. High-angle annular dark-field-

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scanning TEM micrographs of (d) EPS and (e) BSA stabilized CheSeNPs and (f) bright-

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field TEM micrograph of unstabilized CheSeNPs (no capping agent). EDX spectra

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obtained from (g) a single spherical EPS capped CheSeNp, (h) an ensemble of

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spherical BSA capped CheSeNPs and (i) from CheSeNPs nanowires prepared in the

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absence of EPS or BSA. Note that the Cu signal in the EDX spectra is caused by the

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carbon-coated copper support grid used for the TEM analysis.

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Comparison of the capping agents on BioSeNPs, EPS capped CheSeNPs and

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BSA capped CheSeNPs

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FT-IR analysis

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FT-IR spectroscopy provided information of the functional groups present on the

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BioSeNPs (Figure 3, Table S1 in SI). The BioSeNPs had a broad feature between 3404

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to 3270 cm−1, corresponding to -OH and -NH stretching vibrations of amine and

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carboxylic groups.36 Additionally, small but sharp features at 2959, 2928 and 2866 cm−1

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are observed which can be attributed to aliphatic saturated C-H stretching modes.31 The

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strongest feature is observed at 1646 cm−1 and mainly represents the stretching

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vibration of C=O present in proteins (amide I).36 Thus, the feature at 1542 cm−1

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corresponds to the N-H bending vibration in amide linkage of proteins (amide II).31 The

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band around 1460 cm−1 might correspond to methyl groups and/or carboxylate groups

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(antisymmetric stretching vibration), whereas the feature at 1394 cm−1 is most likely

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attributed to the symmetric stretching vibration.37 The presence of carboxylic groups is


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also evidenced by the weak shoulder observed around 1720 cm−1. C-N stretching and

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N-H bending vibrations might contribute to the band observed at 1242 cm−1 (amide-III).31

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The small band at 1151 cm−1 is characteristic for P=O stretching modes.31 Another

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broad feature was observed between 1073 and 1038 cm−1, which corresponds to C-O-C

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and C-H stretching arising from the carbohydrate groups.31,37 The presence of S-H or S-

298

O groups cannot be fully ascertained in the IR spectra due to their weak intensities or

299

due to overlapping bands, respectively.

300 301

Figure 3. IR spectra of (a) EPS stabilized CheSeNPs, (b) extracted EPS, (c) BioSeNPs,

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(d) BSA stabilized CheSeNPs and (e) BSA. Indicated values are in cm−1.

303 304

FT-IR spectra for BioSeNPs, EPS and EPS capped CheSeNPs were similar (Figure 3)

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and confirmed the presence of proteins (amide I: 1653-1646 cm−1, amide II: 1537-1542

306

cm−1 and amide III: 1236-1242 cm−1) and carbohydrates (1040-1077 cm−1) (Fig. 3a-c).

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In contrast, the absence of carbohydrates for BSA and BSA stabilized CheSeNPs is


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shown in the IR spectra by the missing bands in the lower frequency range (Fig. 3d,e).

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The absence of carbohydrates residues is obviously reflected in the OH-stretching

310

region (~3400 cm−1) where reduced band intensities are observed in these spectra31.

311 312

The overall shape of the FT-IR spectra of EPS capped CheSeNPs and EPS was very

313

similar (Figure 3 a,b). Small spectral deviations were observed at 3400 cm−1 for EPS

314

which were shifted to 3420 cm−1 for EPS capped CheSeNPs. The relative intensities of

315

the features around 1450 and 1400 cm−1 are more different for EPS capped CheSeNPs

316

than for EPS. This indicates a change of the carboxylate functional groups due to the

317

presence of the selenium nanaoparticles. Interestingly, the same spectral feature was

318

also observed for BSA stabilized CheSeNPs and BSA (Fig. 3 d,e).

319 320

ζ-potential and HDD measurements

321

The ζ-potential of the BioSeNPs was −41.6 ± 0.5 mV at pH 7.0 and 1 mM NaCl

322

concentration. At the same pH, but with 10 and 100 mM NaCl, the ζ-potential of the

323

BioSeNPs was −29.7 ± 0.7 mV and −17.5 ± 0.9 mV, respectively. Similar negative ζ-

324

potential values were reported for BioSeNPs formed at ambient temperature by

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bacterial cultures, i.e. Bacillus selenatarsenatis and Bacillus cereus.10,38 At a 10 and 100

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mM background NaCl concentration, the iso-electric point of studied BioSeNPs was,

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respectively, 3.2 ± 0.1 mV and 2.4 ± 0.1 mV (Figure 4a, b).


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328 329

Figure 4. ζ-potential measurement of BioSeNP (□), EPS (◊), EPS capped CheSeNPs

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(∆), BSA (○) and BSA capped CheSeNPs (＊) versus pH at (a) 10 mM and (b) 100 mM

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NaCl background electrolyte concentrations. (c) Hydrodynamic diameter of BioSeNP

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(□), EPS capped CheSeNPs (∆) and BSA capped CheSeNPs (＊) versus pH at 100 mM

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NaCl background electrolyte concentration.

334 335

The ζ-potential of EPS, EPS capped CheSeNPs, BSA and BSA capped CheSeNPs

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was, respectively, −35 ± 1.1 mV, −31.5 ± 0.8 mV, −42.9 ± 3.2 mV and −38 ± 0.4 mV at 1

337

mM NaCl concentration and neutral pH. The ζ-potential versus pH curve showed an iso-

338

electric point at pH 2.3 ± 0.1 mV for both EPS and EPS capped CheSeNPs at 10 and

339

100 mM NaCl background electrolyte concentrations (Figure 4a, b). It was not possible

340

to determine the ζ-potential versus pH curve of uncapped CheSeNPs as a small

341

contamination of glutathione affected the ζ-potential versus pH profile. In this


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experiment, the profile of BSA and BSA capped CheSeNPs are a positive control as

343

BSA is known to stabilize the selenium nanoparticles14,24 with a reported iso-electric

344

point of 4.6 ± 0.1 mV39 (Figure 4a, b).

345 346

The ζ-potential versus pH profiles of BioSeNPs, EPS, EPS capped CheSeNPs, BSA

347

and BSA capped CheSeNPs are similar from pH 9.5 to 6.0. However, at pH values

348

below 6.0, BioSeNPs, EPS and EPS capped CheSeNPs follow similar profiles and

349

remain more negative than BSA and BSA capped CheSeNPs (Figure 4). This leads to a

350

similar iso-electric point of BioSeNPs, EPS and EPS capped CheSeNPs (~pHIEP 2.4 at

351

100 mM NaCl concentration) as compared to 4.6 observed for BSA and BSA capped

352

CheSeNPs. It is important to note that the iso-electric point of BioSeNPs (pHIEP 3.2 ±

353

0.1) at 10 mM NaCl background electrolyte was slightly different than the iso-electric

354

point (pHIEP 2.4 ± 0.1) of BioSeNPs at 100 mM NaCl background electrolyte. This

355

change is not significant enough to unambiguously conclude that the differences are

356

due to the interfering background electrolyte.

357 358

It has been shown that the ζ-potential of BioSeNPs loaded with Zn becomes less

359

negative leading to a lowering of their colloidal stability.26 Similar experiments with BSA

360

and EPS capped CheSeNPs suggested that loading of Zn on these CheSeNPs also

361

lead to a less negative ζ-potential of −7.3 and −11.0 mV for, respectively, BSA and EPS

362

capped CheSeNPs contacted with 1000 mg L−1 Zn (Figure S5 in SI). The equilibrium pH

363

varied between 5.5 and 6.5. The zinc concentration required to achieve −5 to −10 mV

364

for BSA and EPS capped CheSeNPs was 10 times more than that required for


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BioSeNPs, even when the concentration of selenium concentration was 4 times higher

366

in the BioSeNPs26. This might be due the smaller size of BSA and EPS capped

367

CheSeNPs (30-50 d-nm) as compared to BioSeNPs (180 d-nm)26.

368 369

The HDD of BioSeNPs increased slightly from 403 ± 8 to 531 ± 6 d-nm when the pH

370

changed from 10.2 to 5.8 at 100 mM NaCl concentration. However, when the pH

371

dropped to 4.7 and 3.8, the HDD increased to 862 ± 29 and 2130 ± 180 d-nm,

372

respectively (Figure 4c). The HDD of EPS capped CheSeNPs and BSA capped

373

CheSeNPs also increased as the pH decreased (Figure 4c). There is a large jump in the

374

HDD of BSA capped CheSeNPs as the pH approached the iso-electric point of BSA. A

375

similar, but smaller jump, in HDD of EPS capped CheSeNPs is observed as the pH

376

approaches the iso-electric point of EPS. HDD measurements at 10 mM NaCl

377

background electrolyte (Figure S6 in SI) gave similar profiles as observed in Figure 4c.

378

It is important to note that it was not possible to compare the HDD versus pH profile of

379

CheSeNPs free of capping agents, as these CheSeNPs had a different shape (wire

380

versus sphere) and they sediment as compared to the capped CheSeNPs (Figure 2).

381 382

DISCUSSION

383

EPS are present on the BioSeNPs

384

This study suggested, for the first time, that the organic layer present on the BioSeNPs

385

synthesized by anaerobic granules is the EPS. The presence of the various functional

386

groups on the BioSeNPs' surface (Figure 1, 3) is due to the attached organic polymers,

387

most likely produced by the microorganisms present in the anaerobic granules. The


19


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presence of carbohydrates, proteins and humic-like substances, which were released

389

upon sonication of the purified BioSeNPs, suggests that EPS comprising of these

390

components is present on the surface of the BioSeNPs. The presence of EPS was

391

further suggested by the presence of amide-I and amide-II bands (proteins) and strong

392

bands at 1073 and 1038 cm−1 (carbohydrates) in the IR spectra (Figure 3, Table S1 in

393

SI). The presence of these carbohydrates, which are always a part of EPS and are not

394

observed in IR spectra of pure proteins31,40, indicates that not protein but EPS

395

containing both proteins and carbohydrates, are present on the BioSeNPs (Figure 3,

396

Table S1 in SI). Moreover, the similar overall shape of the IR spectra and ζ-potential

397

versus pH variation of BioSeNPs, EPS and EPS capped CheSeNPs (Figures 3 and 4a,

398

b) further confirms the presence of EPS on the surface of BioSeNPs. The small DNA

399

concentration found on the surface of BioSeNPs rules out the possibility of cell lysis,

400

thus further confirming the layer on BioSeNPs is from EPS and not due to intracellular

401

organics from the microbial cells. This is an interesting finding as previously only the

402

presence of proteins has been reported on the surface of BioSeNPs15,16 and the origin

403

of the organic layer on the BioSeNPs was unknown17.

404

The similar overall shape of the amide-I and –II modes in IR spectra of EPS and EPS

405

capped CheSeNPs suggests (Figure 3) that the EPS was attached to elemental

406

selenium without major modification in its secondary structure. The distinct shift of some

407

spectral features, e.g. shifting of features from 3400 to 3420 cm−1 and 1384 to 1405

408

cm−1 in EPS capped CheSeNPs as compared to EPS cannot be unambiguously

409

attributed to the interaction of hydroxyl or carboxylic acid groups37 with elemental

410

selenium, respectively, although such assignments appear to be obvious. For instance,


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a slightly different water content in the EPS capped CheSeNPs and EPS samples might

412

cause similar shifts in the spectrum of a KBr pellet used in the sample preparation for

413

FT-IR analysis. However, the band at 1452 cm−1 observed in the spectrum of EPS, but

414

not in the EPS capped CheSeNPs spectrum, most likely represents the antisymmetric

415

stretching mode of carboxylate groups. Thus, the disappearance of this mode reflects

416

the interaction of elemental selenium with these functional groups of EPS. The

417

interaction of EPS and elemental selenium with hydroxyl groups suggests the likely

418

interaction of the carbohydrate fraction of the EPS with elemental selenium, as the

419

sugar residues can be expected to show much more -OH groups. However, further

420

research is required to identify the exact EPS fractions interacting with the CheSeNPs

421

and BioSeNPs.

422 423

The synthesis of BioSeNPs involves two steps: (1) the reduction of selenite to elemental

424

selenium and (2) the subsequent growth of elemental selenium to 50-250 d-nm

425

BioSeNPs with a median of 180 d-nm26 (Figure 5). Reduction of selenite to elemental

426

selenium is carried out intracellularly41, in the periplasmic space22 or extracellulary23 in

427

different microorganisms and thus, the growth of BioSeNPs can take place either

428

intracellularly or extracellularly. It should be noted that this study did not distinguish

429

between the BioSeNPs that grew intracellularly, which are then subsequently expelled

430

outside in the supernatant from those which grew extracellularly. For the elemental

431

selenium that was formed extracellularly, the growth to BioSeNPs takes place in the

432

presence of EPS which is per definition present extracellularly. The elemental selenium,

433

which was formed intracellularly, will most likely have a coating of predominantly


21


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proteins that are involved in their production and subsequent secretion.41 These

435

intracellular elemental selenium particles may partially grow to BioSeNPs coated with

436

proteins.41 The size of these BioSeNPs secreted from the cell is likely to be much

437

smaller (considering the BioSeNPs transport through the cell without damaging the

438

membrane or cell wall). Since the average size of BioSeNPs observed in the majority of

439

the studies is greater than 150 d-nm9,17, further growth of the excreted BioSeNPs can

440

then take place extracellularly, in the presence of the EPS. For the BioSeNPs produced

441

and growing intracellularly but expelled outside the cell due to cell lysis, the full growth

442

would have taken place in absence of EPS and the BioSeNPs are thus capped by

443

predominantly proteins. The smaller quantity of DNA on the BioSeNPs suggests that

444

cell lysis in this study was minimal, thus, the majority of the BioSeNPs observed in this

445

study were formed or growing extracellularly in the presence of EPS.

446

447 448

Figure 5. Scheme demonstrating the growth of elemental selenium to BioSeNPs.

449 450

EPS govern the surface charge of BioSeNPs


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The ζ-potential versus pH curves and iso-electric points of BioSeNPs, EPS and EPS

452

capped CheSeNPs (Figure 4a, b) were similar, while also those of BSA and BSA

453

capped CheSeNPs were identical (Figure 4a, b). This shows that the ζ-potential and

454

iso-electric point of BioSeNPs and EPS capped CheSeNPs are governed by the surface

455

charge of the EPS, rather than that of the elemental selenium. The almost identical iso-

456

electric points of BioSeNPs, EPS and EPS capped CheSeNPs (Figure 4b) further

457

suggests that the EPS is capping the BioSeNPs and CheSeNPs for the entire pH range

458

tested.

459 460

Capping agents can provide the colloidal stability to nanoparticles either by electrostatic,

461

steric or electrosteric mechanisms.12 The sudden jump in HDD of BioSeNPs and EPS

462

capped CheSeNPs (Figure 4c and Figure S6 in SI) at a pH close to the iso-electric point

463

indicates agglomeration of nanoparticles which is a hint for electrostatic stabilization by

464

EPS. The dependence of the ζ-potential on the ionic strength further suggests that the

465

EPS stabilize the BioSeNPs and CheSeNPs by electrostatic repulsion (Figure 4a and

466

b).12 However, due to the bulky structure of EPS, partial stabilization of EPS capped

467

CheSeNPs by steric hindrance cannot be excluded, as demonstrated for the

468

stabilization of silica nanoparticles and quantum dots by BSA.13,42 Thus, this study

469

strongly suggests that the EPS stabilizes the BioSeNPs and CheSeNPs both

470

electrostatically and sterically, thus electrosterically. It is important to note that the larger

471

HDD of BioSeNPs as compared to EPS capped CheSeNPs at a pH exceeding 6 might

472

be due to suboptimum EPS to elemental selenium ratio during the formation of the

473

BioSeNPs.


23


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Manuscript submitted to Environmental Science & Technology 474 475

The carboxylic acid group, whose presence on the BioSeNPs' surface was confirmed by

476

the IR and acid-base titration data, has a pKa value of 3.9 (Figure 1, 3 and Table S1 in

477

SI). Due to their low pKa values, this group will be largely deprotonated at pH values

478

above 5.5 (more than 98%, calculated using the Henderson-Hasselbalch equation). The

479

negative ζ-potential value of the BioSeNPs at a pH below 5.5 suggests a large number

480

of carboxylic acid group sites in the EPS capping the BioSeNPs (Figure 1 and 4).

481 482

Environmental implications

483

This study has demonstrated that EPS present in anaerobic granular sludge are

484

capping the BioSeNPs, govern their surface charge and thus, affect their colloidal

485

properties in the engineered settings where they are synthesized. It is important to point

486

out that the intracellularly grown BioSeNPs will be capped by (a mixture of) proteins and

487

thus their ζ-potential versus pH profile and iso-electric point might be different than that

488

of the BioSeNPs growing extracellularly in the presence of EPS (Figure 4a, b and 5).

489

This would lead to a different colloidal behavior of BioSeNPs that are capped with

490

proteins and those that are capped with EPS in various environmental conditions (pH

491

and interactions with heavy metals, Figure S5 in SI), thus, affecting their fate in the

492

environment.

493 494

The presence of EPS on the surface of BioSeNPs makes them stable in the colloidal

495

suspension and thus, mobile in the low ionic strength and neutral pH environment. This

496

is in contrast to our understanding that EPS of biofilms restrict the dispersion of natural


24

Page 25 of 29



and engineered nanoparticles, as shown for Se43, CdSe, Ag or ZnS nanoparticles.44

498

Thus, this study highlights the importance of further studies on the role of EPS in the

499

fate of bioreduced products of redox active elements in both natural and engineered

500

settings.

501


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ASSOCIATED CONTENT

503

The SI contains methods related to the analytical measurements and extra

504

figures with experimental data as noted in the text. This material is available free of

505

charge via the internet at http://pubs.acs.org

506 507

ACKNOWLEDGMENTS

508

The authors are thankful to Dr. Graciella Gil-Gonzalez (KAUST, Saudi Arabia) for

509

insightful discussion, Ferdi Battles (UNESCO-IHE, The Netherlands) for the Nano-sizer

510

and acid-base titration experiments, Berend Lolkema (UNESCO-IHE, The Netherlands)

511

for TOC, TN and 3D EEM analysis and Elfi Christalle (Helmholtz-Zentrum Dresden-

512

Rossendorf, Germany) for SEM-EDXS measurements and Purvi Jain (Utrecht

513

University, The Netherlands) for DNA measurements.

514

515 516

517

Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding Sources

518

This research was supported through the Erasmus Mundus Joint Doctorate

519

Environmental Technologies for Contaminated Solids, Soils, and Sediments (ETeCoS3)

520

(FPA n°2010-0009).

521


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29

WITHDRAWN: Fate of extracellular polymeric substances during flocculation of anaerobically digested sludge using biogenic flocculant.

Extracellular polymeric substances govern the development of biofilm and mass transfer of polycyclic aromatic hydrocarbons for improved biodegradation.

Extracellular polymeric substances (EPS) of freshwater biofilms stabilize and modify CeO2 and Ag nanoparticles.

Bacterial cell surface properties: role of loosely bound extracellular polymeric substances (LB-EPS).

Biogenic selenium nanoparticles: current status and future prospects.

Biosorption of Hg(II) onto goethite with extracellular polymeric substances.

Optimization for extracellular polymeric substances extraction of microbial aggregates.

Extraction of Structural Extracellular Polymeric Substances from Aerobic Granular Sludge.

Microbial Extracellular Polymeric Substances (EPSs) in Ocean Systems.

Effect of CuO nanoparticles on the production and composition of extracellular polymeric substances and physicochemical stability of activated sludge flocs.

Immobilization of Polymeric Luminophor on Nanoparticles Surface.

Extracellular polymeric substances of a marine bacterium mediated synthesis of CdS nanoparticles for removal of cadmium from aqueous solution.

Electrocoagulation of colloidal biogenic selenium.

Extracellular polymeric substances are transient media for microbial extracellular electron transfer.

Effect of extracellular polymeric substances on the mechanical properties of Rhodococcus.

Characterization of the proton binding sites of extracellular polymeric substances in an anaerobic membrane bioreactor.

Contribution of stratified extracellular polymeric substances to the gel-like and fractal structures of activated sludge.

Influences of extracellular polymeric substances on the dewaterability of sewage sludge during bioleaching.

Isolation of Extracellular Polymeric Substances from Biofilms of the Thermoacidophilic Archaeon Sulfolobus acidocaldarius.

Effect of extracellular polymeric substances on corrosion of cast iron in the reclaimed wastewater.

Scolicidal effects of biogenic selenium nanoparticles against protoscolices of hydatid cysts.

Tannic acid-mediated surface functionalization of polymeric nanoparticles.

Selenopeptides and elemental selenium in Thunbergia alata after exposure to selenite: quantification method for elemental selenium.

Biogenic synthesis of selenium nanoparticles and their effect on As(III)-induced toxicity on human lymphocytes.