Linking composition of extracellular polymeric substances (EPS) to the physical structure and hydraulic resistance of membrane biofilms

Accepted Manuscript Linking composition of extracellular polymeric substances (EPS) to the physical structure and hydraulic resistance of membrane biofilms Peter Desmond, James P. Best, Eberhard Morgenroth, Nicolas Derlon PII:

S0043-1354(17)31053-9

DOI:

10.1016/j.watres.2017.12.058

Reference:

WR 13456

To appear in:

Water Research

Received Date: 15 September 2017 Revised Date:

22 December 2017

Accepted Date: 22 December 2017

Please cite this article as: Desmond, P., Best, J.P., Morgenroth, E., Derlon, N., Linking composition of extracellular polymeric substances (EPS) to the physical structure and hydraulic resistance of membrane biofilms, Water Research (2018), doi: 10.1016/j.watres.2017.12.058. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Graphical Abstract

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Linking composition of extracellular polymeric substances (EPS) to the physical

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structure and hydraulic resistance of membrane biofilms.

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Peter Desmonda,b, James P. Bestc, Eberhard Morgenrotha,b, Nicolas Derlona

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a Eawag, Swiss Federal Institute of Aquatic Science and Technology, 8600 Dübendorf,

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Switzerland

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b ETH Zürich, Institute of Environmental Engineering, 8093 Zürich, Switzerland

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Empa - Swiss Federal Institute for Material Science and Technology, Laboratory for

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Mechanics of Materials and Nanostructures, Feuerwerkerstrasse 39

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CH-3602 Thun, Switzerland

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Email contact of authors: [email protected], [email protected],

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[email protected], [email protected]

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Corresponding author: [email protected]

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First submitted as WR36404 for publication in Water Research on 03.08.2016

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Revised submission on 18.11.2016

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Revised manuscript submitted as WR41267 (formally a new submission) on 15.09.2017

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Revised manuscript submitted as WR41267R2 on 21.12.2017

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Abstract The effect of extracellular polymeric substances (EPS) on the meso-scale physical structure

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and hydraulic resistance of membrane biofilms during gravity driven membrane (GDM)

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filtration was investigated. Biofilms were developed on the surface of ultrafiltration

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membranes during dead-end filtration at ultra-low pressure (70 mbar). Biofilm EPS

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composition (total protein, polysaccharide and eDNA) was manipulated by growing biofilms

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under contrasting nutrient conditions. Nutrient conditions consisted of (i) a nutrient enriched

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condition with a nutrient ratio of 100:30:10 (C: N: P), (ii) a phosphorus limitation (C: N: P

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ratio: 100:30:0), and (iii) a nitrogen limitation (C: N: P ratio: 100:0:10). The structure of the

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biofilm was characterized at meso-scale using Optical Coherence Tomography (OCT).

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Biofilm composition was analysed with respect to total organic carbon, total cellular mass and

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extracellular concentrations of proteins, polysaccharides, and eDNA. 2D- confocal Raman

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mapping was used to characterise the functional group composition and micro-scale

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distribution of the biofilms EPS. Our study reveals that the composition of the EPS matrix can

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determine the meso-scale physical structure of membrane biofilms and in turn its hydraulic

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resistance. Biofilms grown under P limiting conditions were characterised by dense and

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homogeneous physical structures with high concentrations of polysaccharides and eDNA.

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Biofilm grown under nutrient enriched or N limiting conditions were characterised by

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heterogeneous physical structures with lower concentrations of polysaccharides and eDNA.

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For P limiting biofilms, 2D-confocal Raman microscopy revealed a homogeneous spatial

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distribution of anionic functional groups in homogeneous biofilm structures with higher

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polysaccharide and eDNA concentrations. This study links EPS composition, physical

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structure and hydraulic resistance of membrane biofilms, with practical relevance for the

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hydraulic performances of GDM ultrafiltration.

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Keywords: Gravity driven membrane filtration, GDM, Ultrafiltration, Biofilm, Hydraulic

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resistance, Physical Structure, EPS

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1

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The microbial biofilm is akin to a semi-permeable polymeric gel (Wilking et al. 2011). The

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microbial cells embedded in their surrounding matrix of extracellular polymeric substances

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(EPS) collectively form a semi-solid structure which can impede perpendicular flow during

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membrane ultrafiltration (Vrouwenvelder et al. 2016). From an engineering perspective, the

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hydraulic resistance imposed by the biofilm is problematic for membrane filtration systems,

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causing a decline in permeate flux production. Gravity Driven Membrane (GDM)

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ultrafiltration however tolerates biofilm development on the membrane surface for the benefit

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of minimal maintenance, stable flux, and improved permeate quality (Peter-Varbanets et al.

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2010, Peter-Varbanets et al. 2011). Stable but low permeate flux values are reported in GDM

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filters depending of the type of feed water (5 - 15 L/m2/h) (Peter-Varbanets et al. 2010). Thus,

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for successful implementation, reducing the hydraulic resistance of the biofilm for effective flux

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recovery, whilst simultaneously tolerating its presence for stable flux production will be

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imperative. Towards this goal, a fundamental understanding of the influencing factors that

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govern the hydraulic resistance of the biofilms is required for efficient hydraulic operation of

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GDM filtration systems.

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Extracellular polymeric substances (EPS) are a proposed determinant of biofilm hydraulic

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resistance (Dreszer et al. 2013). Relatively little hydraulic resistance can be attributed to the

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embedded microbial cells or inorganic particles (Dreszer et al. 2013, Derlon et al. 2016). A

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comparison of hydraulic resistance profiles from intact biofilms, bacterial cake layers and

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model fouling layers revealed that fouling layers with a greater EPS content had a greater

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hydraulic resistance (Derlon et al. 2016). However, it is not known how the biofilms EPS and

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Introduction

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ACCEPTED MANUSCRIPT their associated physical structures can directly influence hydraulic transport through the

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biofilm (Dreszer et al. 2014, Valladares Linares et al. 2015).

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Dreszer et al. (2013) suggested EPS can limit convective transport through biofilms by

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providing friction to the permeating water molecules. Valladares Linares et al. (2015) support

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this claim when compacting the biofilms physical structure and evaluating the compacted

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biofilms hydraulic resistance. Compaction is proposed to increase the density of EPS, which

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could lead to higher hydraulic resistances on the assumption a higher EPS density can reduce

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the volume between neighbouring EPS molecules, limiting hydraulic flow (Valladares Linares

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et al. 2015). Vrouwenvelder et al. (2016) united such observations in a theoretical model based

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on the Hagen- Poiseuille law which describes hydraulic transport through a channel or pipe. The

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hydraulic resistance imposed by the biofilm is thought to be dependent on the biofilms internal

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architecture, specifically the geometry between the strands of EPS (Vrouwenvelder et al. 2016).

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This theory assumes that volume between the EPS molecules can act as conduits capable of

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hydraulic transport during forced water passage. Numerically it was demonstrated that reducing

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the diameter between EPS molecules can restrict water passage through the channel thus

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increasing resistance to flow during forced water passage (Vrouwenvelder et al. 2016). This is

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in fact analogous to the Darcian models of Tiller and Kwon (1998), who describe fluid flow

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through a cake layer of particles. Therein the authors assume uniform particle size and charge.

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Unlike a layer of inert particles, biofilm EPS is composed of proteins, polysaccharides,

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extracellular DNA (eDNA) and to a lesser extent lipids (Flemming and Wingender 2010). The

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secretion of specific EPS classes is self-regulated based on environmental stimuli (shear,

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nutrient status) (Flemming and Wingender 2010, Danhorn et al. 2004, Vrouwenvelder et al.

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2010). Such biopolymers are capable of electrostatic and functional interactions. In pure-

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culture studies, changes to the composition of EPS can alter the biofilms physical structure

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(Ma et al. 2006, Mayer et al. 1999). It is thus plausible that specific compositions of EPS

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could influence the physical structure of membrane biofilms, e.g., its internal architecture (pores/channels) and in turn permeation through such structures. During membrane filtration,

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the production of extracellular polysaccharides by the biofilm is associated with a lower flux

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production (Chang and Halverson 2003, Chen and Stewart 2002, Herzberg et al. 2009).

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However, whether the production of specific EPS molecules by membrane biofilm leads to

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the formation of specific physical structures with preferential flow paths remains unknown.

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The uncertainty of whether specific EPS molecules can assemble into distinct physical

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structures, which may help or hinder hydraulic transport, is compounded by the fact that

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analysis of physical structures usually occurs at the micro-scale via confocal laser scanning

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microscopy (CLSM). It is argued that CLSM reduces the overall detail of the biofilms

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physical structure and related functions (e.g., permeability) due its measurement range in

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micrometres (Wagner et al. 2010). Furthermore, application of CLSM used in conjunction

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with fluorophore staining (e.g., Concanavalin A) for chemical characterisation (e.g.,

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Polysaccharides) is an invasive method which can alter the physical structure of the biofilm

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(e.g., dehydration, bleaching, decompression) (Neu et al. 2001). For structural observation

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above the micro-scale, Optical coherence tomography (OCT) provides image acquisition in

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the range of millimetres (mm) at the meso-scale (Wagner et al. 2010, Morgenroth and

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Milferstedt 2009). Image acquisition at the meso-scale can provide details of important

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“structure/function” relationships, such as the influence of overall thickness, internal porosity

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and surface topology (roughness) on water passage through the biofilm (Wagner et al. 2010,

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Martin et al. 2014, Fortunato et al. 2017). For in-situ chemical characterisation, 2D-confocal

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Raman microscopy is a non-invasive technique used for characterisation of functional groups

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and their distribution within a sample (Raman 1930, Ivleva et al. 2009, Sijtsema et al. 1998).

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Raman microscopy has been demonstrated to provide accurate chemical information of intact

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microbial aggregates (e.g., flocs). The utility of 2D-confocal Raman mapping in providing

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ACCEPTED MANUSCRIPT localised chemical analysis of EPS in an intact membrane biofilm is unknown (Ivleva et al.

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2009, Blackwell 1977, Wagner et al. 2009). By relating the accumulation of specific EPS

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composition to the meso-scale physical structure of the biofilm, one may better ascertain how

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the chemical composition of the EPS can influence the physical structure and ultimately the

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hydraulic resistance of the biofilm.

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The goal of this study is to understand (i) whether different compositions of the EPS in terms

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of protein, polysaccharide and eDNA concentration can influence the biofilm physical

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structure and (ii) how resulting structures influence the hydraulic resistance of a biofilm

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during GDM ultrafiltration. We hypothesized the hydraulic resistance of the biofilm is

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dependent on the chemical composition of the EPS whereby differences in EPS molecules will

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lead to the formation of distinct physical structures, which have different hydraulic resistances.

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To investigate this hypothesis, different biofilms were grown during GDM filtration on

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membrane surface. Limiting nutrient conditions (N- and P-limitations) were used to produce

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biofilms of different EPS concentrations and compositions (Danhorn et al. 2004, Chen and

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Stewart 2002). OCT analysis was performed to study the effect of EPS accumulation on

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biofilms meso-scale physical structure and ultimately on the biofilms hydraulic resistance.

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The biofilm composition was analysed with respect to total mass, extracellular

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polysaccharide, protein and eDNA concentrations. Results from colorimetric assays were

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confirmed by conducting 2D confocal Raman microscopy characterisations of the different

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types of biofilms.

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2

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2.1

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Membrane fouling simulators (MFS) were operated as GDM membrane filtration systems in

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dead-end mode at a constant transmembrane pressure of 70 mbar (Fig. 1). For each MFS,

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Material and Methods Experimental setup

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ACCEPTED MANUSCRIPT synthetic feed water was pumped into the system with a peristaltic pump (ISM932A, Ismatec,

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Glattbrugg, Switzerland) at a rate ensuring that permeation was solely gravity-driven. The

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system was connected to overhead tanks with overflows through which excess water could

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leave the system. In that way, a constant water head of 70 cm could be maintained. Biofilms

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were grown on ultrafiltration membranes held in the MFS (Eawag, Dübendorf, Switzerland)

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with an area of 18.75 cm2 (Fig. 1).

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[Figure 1]

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2.2

Experimental approach

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Four experimental lines consisting of a negative control (nanopure water), positive growth

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condition (nutrient enriched), a phosphorous limiting and nitrogen limiting conditions were

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run in parallel. Three experimental runs were performed for a duration of 30 days in order to

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assess the reproducibility of our observations. A total of ca. 20-28 flow cells was used per run.

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Duplicate measurements were performed for each time point per each condition and each run,

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i.e., two biofilms characterised after 3, 7, 20 and 30 d of growth. Permeate was collected daily

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for each MFS in 2 L plastic bottles. The experiments were undertaken in an 18 °C

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temperature controlled room.

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2.2.1

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Ultrafiltration membranes (UP150, MicrodyN Nadir, Wiesbaden, Germany) of

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polyethersulfone with a nominal cut-off of 150 kDa were used as a substratum for biofilm

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growth (corresponding to a mean pore size of about 30-40 nm). Membranes coupons were

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cut, rinsed and washed in ethanol to disinfect the membranes surface as described by

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Heffernan et al. (2013). The minimum inhibitory concentration (MIC) required to reduce

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microbial activity of a virgin membrane to that of a nano-pure water was determined to be

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40% (v/v) ethanol diluted in nano-pure water.

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Membrane preparation

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Feed-water preparation

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Sterile feed-water was prepared daily in 10 L glass bottles. All feed-water was autoclaved for

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150 min at 121 °C in a steam sterilizer (Dampfsterilisator LST-V 6-6-9, Belimed, Zug,

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Switzerland or Varioklav HP, Sterico, Wangen, Switzerland) for sterilization. Carbon (C),

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nitrogen (N) and phosphorus (P) were added to nanopure water in the form of acetate

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(CH3COO−), ammonium (NH4+) and ortho-phosphate (PO43−), respectively in a ratio of 100

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(C):30 (N):10 (P) for the positive growth conditions, a ratio representational for drink water.

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A solution of nanopure water with no added nutrients was used as the negative control. P and

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N limitations were imposed to produce a biofilm with distinct EPS composition and

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anticipated different physical structures (Danhorn et al. 2004, Liu et al. 2006, Ras et al. 2011).

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Nutrient limitation was used only as an experimental tool to grow biofilms of different

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physical structures and EPS compositions. Essential trace elements were supplemented to the

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feed-water along with EDTA to ensure trace elements did not precipitate (Hammes and Egli

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2005).

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An overview of the four different nutrient conditions are given in Table 1. Organic carbon

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concentrations are within a range typically expected for drinking water.

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[Table 1]

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2.2.3

Bacteria cultivation and membrane MFS inoculation

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Bacteria for MFS inoculation were cultivated from the Chriesbach river, Dübendorf,

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Switzerland (47°24'16. 3"N 8°36'31. 8"E) during spring time. River water was collected in a

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sterile beaker, sealed and immediately filtered through a 0.45 µm syringe filter to remove

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higher organisms. This was confirmed by visual examination of the filtrate under

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stereomicroscope. 10 mL of filtrate was added to 90 mL of sterile nutrient solution containing

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50 mg C/L and corresponding P and N concentrations under a laminar flow hood. The

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nutrient/filtrate suspension was placed in an orbital incubator for 30 °C, with a rocking speed 9

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synthetic substrate. Bacterial suspension was analysed under stereomicroscope to ensure

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absence of higher organisms after incubation. Bacterial cell number was quantified by ATP

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measurement as described by Hammes et al. (2010): 1.2 · 1010 cells/ml measured. Thereafter,

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the 1.5 mL aliquots were taken and distributed into 2 mL Eppendorf tubes containing 0.5 mL

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of 30% glycerol as a cryo-protectant. Bacterial/glycerol stocks were snap frozen with liquid

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nitrogen and stored at -80 ◦C until use. To regenerate the bacteria from frozen stock, 2 mL of

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the bacteria/glycerol stock was taken and inoculated into 100 mL of sterile nutrient containing

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50 mg C/L with corresponding P and N concentrations in a ratio of 100:30:10 (C: N: P) under

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a laminar flow hood. Suspension was incubated for 18-24 hrs at 30 °C as defined in previous

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steps. For inoculation of the MFS, microbial suspension was centrifuged for 15 min at 5000

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rpm (UniCen MR, Herolab, Germany). The supernatant was decanted allowing resuspension

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of the cell pellet in 30 mL 0.1 M NaCl yielding a final concentration of 1.5 · 1010 cells/ml as

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determined by ATP assay. From the resulting bacterial solution, 2.5 mL was injected into

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each MFS. The spiked MFS was let settle for 18-24 hours to allow filtration of bacterial

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solution allowing known cell coverage over the membrane area. This resulted in a surface

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coverage of 2 · 1013 cells/m2 as determined from ATP assay. Lower ATP activity was noted

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from initial concentration at time of load, likely due to partial loss in cell viability.

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2.2.4

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Permeate flux and hydraulic resistances were derived from the mass of collected permeate.

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The mass of permeate was recorded daily. The volume of permeate was determined by

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dividing the permeate mass by the density of water (1000 kg/m3). The following equations

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were used for the calculations of the permeate flux (J, [L/m2/h])

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Hydraulic parameters

=

∆ ∙∆

(1)

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where ∆V is the change in permeate volume [L], A is the filtration area [m2], ∆t is the change

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time [h]. The total hydraulic resistance (R, [1/m]) was calculated as

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=



(2)

∙

where TMP is the transmembrane pressure [bar], η is the dynamic viscosity of water at a

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given temperature [Pa s], and Rtotal is the total filtration resistance of the fouled membrane

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[1/m]. The biofilm hydraulic resistance (Rbiofilm, [1/m]) was calculated according Eq. (3):

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(3)

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Rbiofilm = Rtotal – Rmembrane

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where Rmembrane is the intrinsic resistance of the clean membrane measured for a period of 24 h

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with nanopure water prior to the bacteria inoculation.

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2.2.5

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Optical Coherence Tomography (OCT) (model 930 nm Spectral Domain, Thorlabs GmbH,

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Dachau, Germany) with a central light source wavelength of 930 nm was used to investigate

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the meso-scale structure of the biofilm. The use of long wavelength light allows to penetrate

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up to a depth of 2.7 mm with axial and lateral resolutions of 4.4 µm and 15 µm, respectively.

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For the image acquisition filtration modules were kept sealed and transferred under pressure

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to the OCT stage. 10-20 B scans (XZ pane image) of 2 × 1 mm or 2 × 1 mm were acquired at

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different time intervals and for 2-3 membrane fouling simulators per each condition. Image

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analysis software developed under Matlab® (MathWorks, Natick, US) was used to analyse

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OCT image. Image analysis consisted of the following steps:

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Structural quantification

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(1) membrane–biofilm interface detection (grey-scale gradient analysis);

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(2) Image binarization (automatic threshold selection);

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(3) calculating physical properties of the biofilm: mean biofilm thickness (z¯ in µm).

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Biofilm harvesting

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Biofilms were removed from the MFS for chemical analysis at each time point. Biofilms were

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scraped from the membrane surface via a cell scrapper and transferred into sterile glass

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beakers. A volume of 30 mL 0.1 M NaCl solution was used to rinse the remaining biofilm

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from the membrane and MFS surface. The collected biomass was re-suspended in the

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washings of the 30 mL 0.1 M NaCl solution. TOC remaining following removal of biomass is

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similar to the one of a virgin membrane, indicating that no left biofilm remained on the

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scrapped membrane. Thus, verifying removal of available biomass from the membranes

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surface.

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2.2.7

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EPS were separated from cells through physical extraction. The approach consisted of

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sonication (homogenisation phase) and centrifugation (separation phase) steps, respectively.

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Sole physical extraction was selected over existing chemical methods to limit contamination

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of sample fractions due to sensitivity of subsequent analytical methods. Different extraction

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tests were performed to maximize EPS extraction while avoiding cell lysis (detected through

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soluble ATP measurement). The harvested biofilm suspensions were placed in an ice bath and

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sonicated for 120 s with 10 s rest intervals using a needle sonicator (Sonoplus HD 3200,

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Bandelin, Berlin, Germany). This was repeated three times to ensure full extraction of the

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biofilms EPS. Thereafter, homogenised samples were centrifuged at 5,000 rpm at 20 ◦C for 15

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min (UniCen MR, Herolab, Germany) to separate the EPS from the cells. Centrifugation was

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repeated to ensure full separation. The supernatant containing the soluble EPS was decanted

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from the resolved cell pellet and aliquots were obtained from the extracellular fraction for

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measurement of total organic carbon (TOC) (TOC-L, Shimadzu, Japan) and EPS analysis

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(Polysaccharide, Protein, eDNA).

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2.2.8 Biofilm density

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Biofilm density (ρbiofilm) was determined as =

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∙



(4)

where M is the biofilm mass (g C), A is the substratum (membrane) surface area (m2), and Lf

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is mean biofilm thickness (m).

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2.2.9 Total polysaccharide quantification

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Concentrations of polysaccharides in the EPS fraction were determined by the Anthrone

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method adapted from Gerhardt et al. (1994). Anthrone was dissolved in absolute ethanol

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before adding it to 75% H2SO4. Absorbance was measured with a UV-Vis spectrophotometer

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(Infinite M200, TECAN, Männedorf, Switzerland) at a wavelength of 625 nm. Glucose was

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used as a standard and results were expressed in mg C/m2 (conversion factor of 0.4 mgC/mg

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glucose).

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2.2.10 Total protein quantification

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Total proteins in the extracted EPS were quantified using a Bicinchoninic Acid (BCA) kit.

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Kits for lower or higher protein concentrations (Micro BCA Protein Assay Kit 23235/ Pierce

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BCA Protein Assay Kit 23225, Thermo Fisher Scientific, Waltham, MA, USA) or for higher

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concentrations were used for the characterization of the different biofilms. The sample to

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reagent ratio was 1:1 for the lower kit and 1:20 for the higher kit. The solutions were

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incubated at 60 °C for 1 h (lower kit), respectively at 65 °C for 30 min (higher kit). After the

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samples were cooled down to room temperature, the absorbance was measured with a UV-Vis

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spectrophotometer (Infinite M200, TECAN, Männedorf, Switzerland) at a wavelength of 562

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nm. Bovine serum antigen (BSA) was used as a standard and results were expressed in mg

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C/m2 (conversion factor of 0.83 mg C/mgBSA).

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ACCEPTED MANUSCRIPT 2.2.11 Extracellular DNA quantification

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Extracellular DNA was determined with the commercially available Quant-iTTM

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PicoGreen® dsDNA Reagent and Kits (Invitrogen/Molecular Probes). The following reagents

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were prepared according to the manufacturer’s instructions. 1 mL of PicoGreen reagent was

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added to 1 mL of sample, 1 x TE buffer or standard in single use cuvettes. Mixtures were

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incubated for 3 min at room temperature. Relative fluorescence was measured at an excitation

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wavelength of 480 nm and an emission wavelength of 520 nm.

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2.2.12 Raman microscopy analysis

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2D Confocal Raman microscopy (model NT-MDT NTEGRA, Spectrum Instruments Ltd.,

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Limerick, Ireland) was performed on an upright confocal Raman microscope with a

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wavelength of 532 nm and a 50X ultra-long working distance objective lens with a numerical

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aperture of 0.55. NTEGRA Spectra provides two separate detection channels: one for

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acquiring the laser confocal (Rayleigh) signal and the second for simultaneous but

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independent collection of the Raman map that reveals the local chemical composition.

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Complete Raman/fluorescence spectrum is recorded in each point of 2D scan. Spectra were

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recorded at a spectral resolution of approximately 2.7 cm−1, with a 2.5 s exposure time for

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each point of the 98 × 98 µm2 Raman map with step size 1.96 µm. Locations (2-3) were

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randomly selected. The laser power was approximately 1 mW. The laser set up was able to

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penetrate through the glass screen of the cell, with the point of focus on the biofilm surface.

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This configuration allowed for effective ‘in situ’ analysis of the biofilms composition.

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Randomized locations were selected for analysis (n = 2-3) and performed on an alternate

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Raman microscope to determine reproducibility.

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2.2.13 Statistical analysis

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Linear regression models were applied to statistically evaluate the effect of growth conditions

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(nutrient enriched, P limiting and N limiting conditions) on the physical and biochemical

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composition of the biofilms (e.g., biofilm thickness, EPS composition). First-order models

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with qualitative variables were used: =

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∙!+

#$

.

∙ &'( )*+. , ∙ ! +

$

.

∙ &'- )*+. , ∙ ! + . ∙ (5)

Where Yt is the response variable (mean biofilm thickness, polysaccharide concentration,

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protein concentration, etc.),

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the slope difference between the enriched-nutrient and N limiting growth conditions and

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.

is the slope under nutrient enriched condition,

#$

is

.

the slope difference between the enriched-nutrient and P limiting growth condition.

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I(N lim.) and I(P lim.) are indicator variables (equal to 0 or 1 depending of the data set that is

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considered). The errors . are assumed to be independent and normally distributed. These

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assumptions allow to test statistically if the parameters are different from zero (the null

325

hypothesis) (Harrell Jr 2015). For example, the slope difference between the enriched and N

326

limiting growth conditions is statistically different from zero if the null hypothesis

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0 is rejected. The model described by Eq. (5) was applied to the entire data set (e.g. TOC n =

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16 MFS) (full data set analysed provided in supplementary information (SI).

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3

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3.1 Permeate flux and hydraulic resistance

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3.1.1

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The change in the mean flux of each experimental condition was monitored for 30 days for 3

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separate experimental runs to determine experimental reproducibility (Fig. 2a-c). Flux

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stabilisation occurred only under nutrient enriched (Fig. 2a) and N-limiting (Fig. 2c)

335

conditions but not for P limiting condition (Fig. 2b) which decreased up to the experimental

336

endpoint. This was standard for all three experimental runs. Under N limiting conditions, flux

337

stabilisation was the higher, producing an average flux production of 10 ± 1.2 L/m2 /h, (n = 12

#$

.

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Permeate flux

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MFS) between day 19-30 compared to an average stable flux of 4.5 ± 2.7 L/m2 /h (n = 12

339

MFS) for the nutrient enriched condition. Permeate flux production for P limiting condition

340

however declined continuously to 1.60 ± 0.7 L/m2 /h (n = 12 MFS) up to 30 days.

341

3.1.2

342

The biofilms hydraulic resistance was derived from mean flux values monitored for 30 days

343

repeated for three separate experimental runs (Fig. 2d-f). Hydraulic resistance for the biofilms

344

formed under the P limiting condition (Fig. 2e) increased over time, reaching 1.26·1012 ±

345

2.07·1011 1/m (n = 12 MFS) after 30 days of growth. Conversely, the hydraulic resistance of

346

biofilms formed under the nutrient enriched condition (Fig. 2d) increased to an average of

347

3.78·1011 ± 2.75·10 (n = 12 MFS) after 10 days of growth and remained stable at an average

348

resistance 3.78·1011 ± 2.75·10 1/m (n = 12 MFS) thereafter until 30 days of system operation.

349

Similarly, the biofilm formed under the N limiting condition (Fig. 2f), had a lower hydraulic

350

resistance of 2.25·1011 1/m after 30 days of growth compared to P limiting and nutrient

351

enriched conditions respectively.

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Biofilm hydraulic resistance

[Figure 2]

352

3.2 Biofilm physical structure

354

3.2.1

355

Visual observations at the meso-scale revealed a time dependent accumulation of biomass on

356

the surface of membranes (Fig. 3). Contrasting growth conditions produced biofilms with

357

different physical structures at meso-scale, associated with different hydraulic resistances.

358

Biofilms grown under nutrient enriched conditions had a heterogeneous morphology (n = 50).

359

Meanwhile, the biofilm of the P limiting growth condition had a largely homogenous

360

morphology (n = 50). The biofilm grown under the N limiting condition formed a thin and

361

heterogeneous structure (n = 30).

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363

meso-scale structure. Biofilms with homogenous morphologies had very high hydraulic

364

resistances. Conversely, heterogeneous biofilm morphologies were associated with

365

significantly lower hydraulic resistances. Biofilms formed under the nutrient enriched

366

condition, while also being heterogeneous, had a higher hydraulic resistance than the biofilm

367

of the N limiting condition.

RI PT

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369

3.2.2

370

Morphological observations at the meso-scale were supported by quantification of mean

371

biofilm thickness based on OCT image analysis as described in section 2.2.5. The thickness of

372

the nutrient enriched biofilm (Fig. 4a) increased over time, achieving an average thickness of

373

803 ± 70 µm (n = 37 locations) after 30 days of growth. The biofilm of the P limiting (Fig.

374

4b) and N limiting (Fig. 4c) conditions were thinner achieving respective thickness of 251 ±

375

18 µm (n = 17 locations) and 214 ± 37 µm (n = 30 locations) after 30 days of growth. The

376

thickness of the enriched nutrient film was significantly higher than the thickness of both the

377

P and N limiting biofilms, with a p-value of 3e-04 and 4e-05 respectively.

378

Increase in thickness of the nutrient enriched biofilm had a limited impact on its hydraulic

379

resistance. The homogenous P limiting biofilm had a higher hydraulic resistance compared to

380

the thicker biofilm of the nutrient enriched despite being thinner. Biofilms formed under the N

381

limiting condition were thinner than the biofilm of the nutrient enriched and had a lower

382

hydraulic resistance indicating passage length may influence the hydraulic resistance of

383

biofilms with similar morphological distributions of biomass.

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3.3 Biofilm accumulation

385

3.3.1

386

During the first 7 days of operation, the nutrient enriched condition (Fig. 4d) yielded an

387

average TOC of 463 ± 15 mg C/m2 (n = 4) compared to 312 ± 8 mg C/m2 (n = 4) and 206 ±

388

1.12 mg C/m2 (n = 5) of P (Fig. 4e) and N limiting conditions (Fig. 4f). As the biofilms

389

matured onwards of 7 days, higher concentrations of biomass in terms of TOC were measured

390

for the P limiting condition, yielding a final concentration of 1,006 ± 32 mg C/m2 (n = 4) after

391

30 days of growth compared to 760 ± 40 mg C/m2 (n = 6) and 400 ± 80 mg C/m2 (n = 4) for

392

enriched and N limiting growth conditions after 30 days of growth respectively. Data analysis

393

indicated that the mass of the enriched-nutrient and P limited biofilms are similar (null

394

hypothesis not rejected, p-value of 0.7), while the mass of the N limited biofilm is

395

significantly lower than the one of the enriched-nutrient biofilms (null hypothesis rejected

396

with a p-value of 0.009).

397

3.3.2

398

As the biofilms matured, the biofilm formed under the P limiting condition had a higher

399

density of 3 ± 0.22 kg C/m3 (n = 2) compared to densities of 0.9 ± 0.198 kg C/m3 and 1.5 ±

400

0.27 kg C/m3 for the nutrient enriched and N limiting conditions respectively after 30 days of

401

growth.

402

3.3.3

403

As biofilms matured, biofilms formed under nutrient enriched conditions had higher active

404

cell mass compared to P and N limiting conditions respectively in all experimental runs.

405

Biofilms formed under nutrient enriched conditions contained 7.9·10-7 ± 5.410-7 moles of

406

ATP/m2 (n = 3) after 30 days of biofilm growth. This corresponded to 3.9·1011 ± 2.8·10 active

407

cells/m2 assuming 2·10-18 moles of ATP per unit cell. Biofilms formed under P and N limiting

408

conditions contained 7 ·10-8 ± 1.9·10-8 moles of ATP/m2 (n = 3) and 2.4·10-8 ± 6.3·10-09 moles

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Biofilm activity

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ACCEPTED MANUSCRIPT of ATP/m2 (n = 3) respectively. This corresponded to 3.5·1010 ± 9.6·109 (n = 3) and 1.2·1010 ±

410

3.1·109 active cells/m2 assuming 2·10-18 moles of ATP per unit cell for biofilms formed under

411

P and N limiting conditions respectively. This indicates lower cellular activity in biofilm

412

grown under nutrient limiting conditions. [Figure 4]

413

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414

3.4 EPS concentration

415

3.4.1

416

Extracellular protein concentration increased in all biofilms during 30 days of growth (Fig. 5).

417

Biofilms formed under nutrient enriched (Fig. 5a) conditions yielded 461 ± 8.2 mg C/m2 (n =

418

5) after 30 days of growth compared to 186 ± 6.37 mg C/m2 (n = 5) and 198 ± 1.67 mg C/m2

419

(n = 2) for P (Fig. 5b) and N limiting (Fig. 5c) conditions respectively. Statistical analysis

420

indicated that the null hypotheses (similar protein concentrations in the P and N limiting

421

biofilms compared to nutrient enriched biofilms) can be rejected with p-values of 0.0002 and

422

2e-05, respectively.

423

3.4.2

424

During early biofilm development (up to 7 days) biofilms formed under the nutrient enriched

425

conditions (Fig. 5d) yielded 188 ± 5 mg C/m2 (n = 4), compared to 306 ± 23 mg C/m2 (n = 4)

426

yielded under P (Fig. 5e) and N limiting (Fig. 5f) conditions respectively. After 30 days of

427

biofilm growth biofilms formed under P limiting conditions (Fig. 5e) yielded 1,006 ± 32.48

428

mg C/m2 (n = 5) compared to 188 ± 2.94 mg C/m2, (n = 4) and 110 ± 30 mg C/m2, (n = 5) for

429

nutrient enriched (Fig. 5d) and N limiting (Fig. 5f) conditions respectively. The accumulation

430

of polysaccharides in the P limiting biofilms was statistically higher than in the nutrient

431

enriched biofilms (null hypothesis, i.e., similar polysaccharide concentrations in the P limiting

432

and enriched biofilms was rejected with p-value of 3e-08). The accumulation of

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ACCEPTED MANUSCRIPT 433

polysaccharides in the N limiting biofilms compared to the nutrient enriched biofilms was

434

also statistically significant (null hypothesis, i.e., similar polysaccharide concentrations in the

435

N limiting and enriched biofilms was rejected with p-value of 0.0034).

436

3.4.3

437

eDNA concentration increased over the course of biofilm growth (Fig. 5). Biofilms formed

438

under P limiting (Fig. 5h) yielded 159 mg eDNA/m2 (n = 2) compared to 82 mg eDNA/m2 (n

439

= 2) and 60 ± 1 mg eDNA/m2 (n = 2) yielded under nutrient enriched (Fig. 5g) and N limiting

440

(Fig. 5i) growth conditions. Change in the eDNA concentrations of the P limited biofilm was

441

statistically different than in the nutrient enriched biofilms (null hypotheses, i.e., similar

442

eDNA concentrations in the P and N limiting biofilms compared to the enriched biofilms

443

were rejected with p-values of 4e-05). The eDNA content of the N limited biofilm was

444

different from the ones of the enriched-nutrient biofilms (p-value of 0.025).

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eDNA concentration

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3.4.4

In-situ Raman microscope analysis of membrane biofilms

447

In-situ Raman microscope was performed to characterise the functional group profile of

448

different biofilms with distinct EPS matrix composition and meso-scale physical structures to

449

better understand the link between polymer composition and biofilm physical structure.

450

Figure 6 illustrates the Raman spectra of nutrient enriched, P and N limiting conditions. For

451

all conditions, the spectra show 3 identical bands with different strengths of intensity

452

depending on the biofilm growth condition at; (i) ~1,000 1/cm; (ii) ~1,155 1/cm; and (iii)

453

~1,510 1/cm. The assignment of Raman bands is based on reference samples for

454

polymicrobial biofilms validated with surrogate substances (Ivleva et al. 2009, Blackwell

455

1977, Wagner et al. 2009). The band at approximately 1,000 1/cm (i) corresponds to O-P2- the

456

negatively charged phosphate backbone of the DNA/RNA molecule (Ivleva et al. 2009,

457

Blackwell 1977). Bands at ~1,155 1/cm (ii) are typical for vibrations of glucosidic ring

AC C

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20

ACCEPTED MANUSCRIPT structures of long change polysaccharides (Wagner et al. 2009, Berg et al. 2002). Shoulder

459

bands between 1,400-1,420 1/cm (iii) are typical for COO- 1/cm (Blackwell 1977), while at

460

1,510 1/cm strong C=C stretching modes are allocated (Ivleva et al. 2009). A C-C stretch is

461

also detectable around 855-899 1/cm and corresponds to a non-compound specific covalent

462

bond between two carbon atoms and can constitute the backbone of a long chain

463

polysaccharide or a peptide compound (Blackwell 1977, Berg et al. 2002).

464

The intensity of the specific Raman shifts is viewed with respect to the base line (back-ground

465

signal) to the maximum of the peak. Biofilms grown under the nutrient enriched and P

466

limiting conditions show higher organic signal intensities at (i) ~1,000 1/cm; (ii) ~1,155 1/cm;

467

and (iii) ~1,510 1/cm – compared to the enriched biofilms and N limiting biofilms. The spatial

468

distribution of bands a, b and c varied in space on the micro-scale for each biofilm grown

469

under different nutrient conditions. Under P limiting conditions, Raman intensity was

470

homogeneous distributed in space compared to nutrient enriched and N limiting conditions.

471

Overall, the 2D Raman microscope confirmed results from colorimetric assays, e.g., the

472

significantly higher polysaccharide content of the biofilms grown under P limitations.

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[Figure 6]

474

4

475

4.1 Distinct biofilm physical structures have a characteristic hydraulic resistance

476

Our study links physical structure of membrane biofilms to hydraulic resistance. Under P

477

limiting conditions, biofilms formed dense, uniform physical structures with low

478

permeability, as indicated by a high hydraulic resistance (Fig. 3). Under the nutrient enriched

479

and N limiting conditions, biofilms were characterised by low density values and in turn by

480

low hydraulic resistances.

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Discussion

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21

ACCEPTED MANUSCRIPT The nutrient enriched biofilm was thicker and had a higher hydraulic resistance compared to

482

biofilms formed under the N limiting condition—despite both having a similar heterogeneous

483

morphology. Thus, it is proposed that the passage length (thickness) can greatly influence the

484

biofilm hydraulic resistance in biofilms with a comparable morphology (heterogeneity). Using

485

Hagen–Poiseuille's law for laminar flow through a biofilm one can evaluate the respective

486

influence of biofilm thickness and, to an extent, porosity on permeation through the biofilm

487

(J) (Vrouwenvelder et al. 2016). Hagen–Poiseuille's law relates the mean velocity through a

488

cylindrical pore to the trans-membrane pressure applied over the length of the pore (TMP),

489

the length (Lf) and the diameter of the pore (dp2) for a given dynamic viscosity (µ). From the

490

Hagen–Poiseuille equation the flux can be calculated considering the total cross-sectional area

491

of the pores (Ap) relative to the total membrane area (Am):

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012∙354 ;4

= 67 ∙ 8 ∙ 9 ·;

492

:

(6)

<

As can be seen from Eq. (6) the flux is related to the inverse of the thickness of the pore but

494

increases with the square of its diameter and total cross-sectional area of the pores per

495

membrane surface. Thus, larger (and more) pores significantly increase the flux and decrease

496

the hydraulic resistance. Conversely, P biofilms had a hydraulic resistance several orders of

497

magnitude higher than the N limiting biofilm, despite having a comparable hydraulic

498

resistance. We suggest the higher hydraulic resistance of biofilms formed under P limiting

499

conditions compared to the nutrient enriched and N limiting condition is due to a lower

500

porosity. This hypothesis is supported by comparison of biofilms density. Biofilms grown

501

under P limiting conditions had a significantly higher density than those formed under

502

nutrient enriched and N limiting conditions respectively. Given passage length of the nutrient

503

enriched biofilm is greater, we conclude that the hydraulic resistance of membrane biofilms

504

has greater dependency on density than biofilm thickness.

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ACCEPTED MANUSCRIPT 4.2 Linking EPS chemical composition to biofilm physical structure

506

For all biofilms, increasing EPS mass positively correlated with increasing hydraulic

507

resistance. This observation is comparable to the findings of Derlon et al. (2016). Therein,

508

cake layer structures with greater concentrations of EPS, rather than cell mass, had a higher

509

hydraulic resistance (Derlon et al. 2016). However, in the current study, P limiting biofilms

510

had a higher hydraulic resistance than nutrient enriched conditions despite having similar

511

concentrations of EPS after 20 days of growth (Fig. 2a, b and Fig. 4e, d).

512

We suggest EPS composition is linked to the physical structure of the biofilm and in turn its

513

hydraulic resistance. EPS produced under P limiting conditions contained statistically higher

514

concentrations of polysaccharides and eDNA and accumulated into dense homogeneous

515

physical structures (Fig. 6b). Homogeneous biofilm structures with high concentrations of

516

polysaccharides and eDNA were characterised by a Raman spectra indicative of C-O-C

517

groups, Carboxyl groups (COO-) and O-P2- (DNA backbone) homogeneously distributed in

518

space (Fig. 6a and c). Such functional groups are molecular finger prints of anionic

519

polysaccharides and DNA biopolymers (Sutherland 2001a, b). We suggest the formation of a

520

homogeneous physical structure is linked to a high content in the functional groups capable of

521

strong intermolecular interaction and their homogeneous spatial distribution. This is based on

522

the fact that the C-O-C groups detected at ~1,155 1/cm are, for example, implicated in

523

glycosidic bond formation, a form of strong covalent interaction which can link

524

polysaccharide molecules to another saccharide groups (e.g., α-1,4-glycosidic bond, 335

525

kJ/mol) (Berg et al. 2002, Hibben 1936). The presence of C-O-C groups is also indicative of

526

nitrogen-carbon linkage between the 9' nitrogen of purine bases and the 1' carbon of the sugar

527

group in DNA (Berg et al. 2002). Carboxyl groups (COO-) detected at ~1,510 1/cm can also

528

provide additional electrostatic interactions between neighbouring negatively charged

529

polymers via a divalent cation intermediary such as Ca2+ (Mayer et al. 1999, Laue et al. 2006).

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23

ACCEPTED MANUSCRIPT The resulting salt bridge can link two negatively charged residues via hydrogen bond

531

formation (10-30 kJ/mol) to form an array of repeating polymeric blocks, termed an “egg-

532

box” configuration (Lewandowski and Beyenal 2013). Such interactions can form uniform

533

packing structures and homogeneous physical structures which are typical of alginate gel

534

layers (Giraudier et al. 2004, O'Neill et al. 1983). Similarly, the negatively charged DNA

535

backbone O-P2- identifiable at 1,095 1/cm can also interact with the divalent inorganics, again,

536

bridging and stabilising neighbouring anionic polymers to form a uniform cross-linked matrix

537

of interacting polymers (Böckelmann et al. 2006, Peterson et al. 2013). In the current study,

538

higher concentrations of polysaccharide and eDNA did, in fact, coincide with a strong signal

539

intensity for carboxylic and O-P2- functional groups which were homogeneously distributed in

540

space, forming a densely packed meso-scale physical structure.

541

Reduced secretion of polysaccharides and eDNA, or their degradation by catabolic enzymatic

542

activity, has also been reported to lead to the formation of dispersed/heterogeneous biofilm

543

structures, albeit in simple biofilm colonies (Ma et al. 2006, Laue et al. 2006, O'Neill et al.

544

1983, Ahimou et al. 2007, Boyd and Chakrabarty 1995). In our study, lower concentrations of

545

polysaccharides and eDNA coincided with greater heterogeneity in functional group

546

distribution and in the formation of a heterogeneous physical structures. We propose the high

547

concentrations of polysaccharides and eDNA favour close range electrostatic interactions

548

leading to the formation of a homogeneous physical structures of low permeability. At lower

549

concentrations, functional groups are heterogeneously distributed in space, limiting potential

550

for electrostatic interactions, imperative for the formation of an otherwise dense homogeneous

551

structure.

552

The scientific implication of the presented work links in-situ, functional group composition

553

and distribution to the formation of a specific meso-scale physical structure, without alteration

554

to its nascent composition or physical structure. This has allowed greater insight into the form

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24

ACCEPTED MANUSCRIPT (e.g., chemical/structural) and function (hydraulic permeability) of microbial biofilms, with

556

practical relevance for the performance of membrane filtration systems.

557

4.3 Practical considerations

558

In the current study, growth conditions in terms of nutrient limitations impacted cellular

559

activity, concentration and composition of extracellular polymeric substances, as well as the

560

biofilm structure and hydraulic resistance. The influence of the feed water composition on

561

biofilm hydraulic resistance has been reported for different types of filtration systems. In the

562

case of bio-filters, nutrient enrichment by low-level phosphorus supplementation can also

563

enhance the hydraulic performance, decreasing head-loss by 15% due to the formation of

564

heterogeneous biofilm structures (Lauderdale et al. 2012). In our study, nutrient enriched

565

conditions allowed the formation of a permeable biofilm with a stable hydraulic resistance.

566

This was linked to the formation of “open” heterogeneous biofilm structures (Fig. 3).

567

In membrane filtration systems, operated under cross-flow conditions, Vrouwenvelder et al.

568

(2010) observed that P limiting regimes can limit biofilm formation over a period of 4 days.

569

Therein, it is suggested P limiting conditions restricts cellular activity and in turn biofilm

570

growth (Vrouwenvelder et al. 2010). The difference between our observations (increased EPS

571

and hydraulic resistance under P limitations) and the observations of Vrouwenvelder et al.

572

(2010), may be attributed to the different time-scales (30 vs. 4 d) or to the different filtration

573

modes (dead-end vs. cross-flow). In our study, during the first 4 days with P limitation

574

conditions, cellular growth, EPS production and hydraulic resistance were in fact lower

575

compared to nutrient enriched conditions. Long term application of P limiting conditions led

576

to greater accumulation of EPS, thereby increasing hydraulic resistance. This is in agreement

577

with previous authors who report P limitation can enhance biofilm formation in mono-species

578

biofilm models (Danhorn et al. 2004, Rüberg et al. 1999). Danhorn et al. (2004) demonstrate

579

P limitation can enhance biofilm formation in mono-species biofilms through the PhoR-PhoB

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555

25

ACCEPTED MANUSCRIPT regulatory system. Exopolysaccharide production by S. meliloti is also regulated through the

581

PhoB response regulator, which activates production of galactoglucan (EPS II) under

582

phosphate limitation by direct activation of the exp gene—in a pathway akin to the bacterial

583

stringent stress response (Rüberg et al. 1999, Boutte and Crosson 2013). For polymicrobial

584

biofilms, exemplified in the current study, the reasons for increased EPS production under P

585

limiting conditions are yet to be investigated. Supporting literature describing the impact of

586

nutrient limitations on biofilm structure could not be found.

587

The practical implication of the presented work demonstrates that restricting microbial

588

activity by P limitation does not stop biofilm accumulation and can lead to its exacerbation

589

due to over production of EPS. This challenges the explanation of Vrouwenvelder et al.

590

(2010), who suggests P limitation can limit biofilm accumulation by reducing cell

591

proliferation and thereby restrict biofouling. We demonstrate P limitation can reduce cell

592

growth, but can simultaneously increase accumulation of EPS responsible for the formation of

593

a dense biofilm structure with a high hydraulic resistance.

594

5

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the extracellular polymeric substances in membrane biofilms and in turn the biofilms

596

physical structure and its hydraulic resistance.

597 598 599

Growth conditions in terms of nutrient limitations determine chemical composition of

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Conclusions

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•

Biofilms with a high polysaccharide and eDNA content (P limiting conditions) formed

dense homogeneous structures with high hydraulic resistance. The biochemical

600

composition of homogeneous biofilm structures was characterised by a high content in

601

functional groups capable of strong covalent and hydrogen bond formation (C-O-C

602

groups, carboxyl groups (COO-) and O-P2- (DNA backbone)).

26

ACCEPTED MANUSCRIPT 603

•

Biofilms with low polysaccharide and eDNA content (nutrient enriched and N limiting conditions) formed thick, heterogeneous structures imperative for low hydraulic

605

resistance. The biochemical composition of those heterogeneous physical structures

606

was characterised by lower abundance of functional groups capable of strong covalent

607

and hydrogen bond formation.

608

•

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604

For practice, reducing microbial activity on the surface of the membrane by nutrient limitation does not necessarily prevent biofilm accumulation. When developing

610

biofilm control measures, understanding the link between biofilm nutrition, biofilm

611

physiology (e.g. EPS production), and hydraulic performance is imperative.

612

613

Supplementary information (SI)

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1. OCT images for Runs 2 and 3 corresponding to Fig. 3 available in SI.

615

2. Fitted data set for the statistical analysis and detailed results from the statistical

EP

617

analysis are provided.

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27

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ACCEPTED MANUSCRIPT Table 1 Composition of nutrient feed-solution: N [mg /L]

P [mg /L]

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-

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Nutrient enriched

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1.5

0.5

P limiting

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1.5

-

N limiting

5

-

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Figure 1 Schematic of membrane fouling simulator used for biofilm growth and hydraulic

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characterization.

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Figure 2: Change in the permeate flux (a-c) and biofilm hydraulic resistance (d-f) of biofilms

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formed under the nutrient enriched, P limiting and N limiting conditions for three spearate

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runs (n = 24, flow cells per condition). Error bars: standard deviation of mean of all flow cells

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per each run.

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Figure 3: OCT images of different biofilm morphologies monitored over 30 days of growth

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under contrasting nutrient conditions. White pixel: Biomass accumulation, Black: Absent/low

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biomass accumulation. Scale bar: 200 µm. Red: Marked membrane. Images for all runs are

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available in supplementary information (SI)

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Figure 4: Mean biofilm thickness (a-c) and mass accumulation (d-f) with respect to TOC

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over 30 days of biofilm growth under contrasting nutrient conditions. Error bars: standard

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deviation of 4-6 MFS, (n = 4-6). Note that thickness and TOC were not measured in all runs.

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Figure 5: Change in the concentrations of extracellular proteins, polysaccharides and eDNA

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over 30 days of biofilm growth under contrasting nutrient conditions. Error bars: standard

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deviation of an average of 2 independent runs consisting of an average 2 MFS per run (n = 4

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MFS). Note that measurements required sacrificial sampling and are not available for all runs.

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Figure 6: 2-D confocal Raman microscopy of intact membrane biofilms formed under

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nutrient enriched, P limiting and N limiting conditions after 30 days of growth. Averaged

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Raman spectra (of a 98 µm2 scanning area) shows three distinct bands (i-iii) of varying

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relative intensity. 2D confocal Raman mapping shows the spatial distribution of the

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corresponding maximum intensity projections (d-f) for nutrient enriched, P limiting and N

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limiting conditions respectively. Scale bar: 10 µm. Spectrum baseline: background

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signal/glass surface.

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Figure 1 Schematic of membrane fouling simulator used for biofilm growth and hydraulic

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characterization.

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Figure 2 Change in the permeate flux (a-c) and biofilm hydraulic resistance (d-f) of biofilms

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formed under the nutrient enriched, P limiting and N limiting conditions for three spearate

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runs (n = 24, flow cells per condition). Error bars: standard deviation of mean of all flow cells

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per each run.

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Figure 3: OCT images of different biofilm morphologies monitored over 30 days of growth

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under contrasting nutrient conditions. White pixel: Biomass accumulation, Black: Absent/low

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biomass accumulation. Scale bar: 200 µm. Red: Marked membrane. Images for all runs are

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available in supplementary information (SI)

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Figure 4: Mean biofilm thickness (a-c) and mass accumulation (d-f) with respect to TOC

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Figure 5: Change in the concentrations of extracellular proteins, polysaccharides and eDNA

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deviation of an average of 2 independent runs consisting of an average 2 MFS per run (n = 4 MFS). Note that measurements required sacrificial sampling and are not available for all runs.

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Figure 6: 2-D confocal Raman microscopy of intact membrane biofilms formed under nutrient enriched, P limiting and N limiting conditions after 30 days of growth. Averaged

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Raman spectra (of a 98 µm2 scanning area) shows three distinct bands (i-iii) of varying relative intensity. 2D confocal Raman mapping shows the spatial distribution of the corresponding maximum intensity projections (d-f) for nutrient enriched, P limiting and N

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Highlights: Hydraulic resistance of biofilms formed during GDM filtration was studied.

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Influent composition determined EPS chemical composition.

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Resulting EPS compositions were linked to distinct physical structures

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Hydraulic resistance was dependent on biofilm thickness and density

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