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Influence of Feed Composition on the Monomeric Structure of Free Bacterial Extracellular Polysaccharides in Anaerobic Digestion Chencheng Le, and David C Stuckey Environ. Sci. Technol., Just Accepted Manuscript • Publication Date (Web): 31 May 2017 Downloaded from http://pubs.acs.org on June 1, 2017

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Environmental Science & Technology

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Influence of Feed Composition on the Monomeric Structure of

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Free Bacterial Extracellular Polysaccharides in Anaerobic

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Digestion

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Chencheng Lea,b, and David C. Stuckeya,c*

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a

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Institute, Nanyang Technological University, 1 Cleantech Loop, CleanTech One, Singapore

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637141, Singapore

Advanced Environmental Biotechnology Center, Nanyang Environment & Water Research

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b

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Environmental Engineering, Nanyang Technological University, 50 Nanyang Avenue,

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Singapore 639798, Singapore

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c

Division of Environmental and Water Resources Engineering, School of Civil and

Department of Chemical Engineering, Imperial College London, SW7 2AZ, U.K.

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*Corresponding author Address: Room 510, ACE Building, South Kensington, LONDON SW7 2AZ; Tel: +44 (0) 207 594 5591; Fax: +44 (0) 207 594 5629; Email address: [email protected];

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ABSTRACT

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Six 5.0-liter fill-and-draw batch reactors were used with different feed compositions

22

containing a range of carbohydrates (glucose, sucrose, fructose) and nitrogen sources (urea,

1 Environment ACS Paragon Plus


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NH4Cl) at various concentrations to investigate free extracellular polysaccharide (EPS)

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production during anaerobic digestion (AD). This work analyzed not only their

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monosaccharide components, but also their specific linkage patterns as well as the change

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associated with different chemical nature in carbon substrates or nitrogen sources; all of these

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parameters can have profound biological consequences, and were correlated to

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macronutrients present in the feed. It is believed that feed composition is a major factor

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which determines the physicochemical characteristics of the free EPS. Our findings also

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suggest that the differences associated with the digestion of various carbon substrates and/or

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nitrogen sources could alter monomeric saccharide composition and concentrations of the

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free EPS. Such insights demonstrate that previous studies on feed C/N ratios tended to

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overestimate EPS production, while variations in the chemical nature of the nitrogen source

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were overlooked. Our results also link the physiochemical properties of free EPS with

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underlying biofouling mechanisms, and demonstrate that membrane fouling is to some extent

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dependent upon the prevailing nutritional environment and feed composition.

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Graphical Abstract

38 39


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1. Introduction

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Free extracellular polysaccharides (EPS) are a complex assortment of extracellular polymeric

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substances secreted by a broad range of bacterial species.1 The free EPS can be found on the

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outermost surface of a wide range of bacteria, but in its unbound form it only maintains a

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limited association with the surface of bacterial cells.2,3 Moreover, certain free EPS used to

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be part of the capsular polysaccharides that may themselves be released into the growth

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medium (i.e. become free) as a consequence of their weak adhesion properties.1,4 The

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common biological functions of bacterial EPS includes resistance to desiccation5, protection

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against harmful substances, and a permeability barrier that facilitates the selective

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transportation of nutrients1. More significantly, EPS plays a major role in mediating the

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bacterial colonization of surfaces by enabling cell adhesion and co-aggregation via dipole

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interactions, covalent or ionic bonding, steric interactions, and hydrophobic association.6-10

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Finally, of recent concern is that membrane autopsies in membrane bioreactors reveal a

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significant amount of fouling resistance is due to an uneven distribution of EPS11, and this

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resistance directly correlated with EPS which undergoes intermolecular or intramolecular

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ionic cross-linking, and subsequently clogs the membrane pores and leads to further

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fouling12,13.

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Unsurprisingly, EPS are as functionally and structurally diverse as the bacteria that

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synthesize them. Since they can be present in many forms, understanding the chemical basis

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for such a physical distinction is of considerable importance. Considerable effort has been

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expended to catalogue the enormous structural complexity of EPS, which is made possible by

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the wide assortment of monosaccharide combinations available, and linkage types, and to

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elucidate their biosynthesis and export. Analysis of polysaccharide structures was termed



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“glycomics”, and has received considerable interest in recent years14, but its progress lags far

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behind proteomics and genomics, partly because of analytical and preparative difficulties.

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Since carbohydrates or polysaccharides cannot be “amplified” like nucleic acids, there is no

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template that encodes for their sequence. Beside the technical challenges associated with their

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unique chemical properties, another significant challenge in analyzing them derives from

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their structural complexity.15,16 Even though there are only nine monosaccharides commonly

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found in biological systems, with three different ways of forming linkages between each

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monomers, a short chain of four monosaccharides can result in 15 million possible

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combinations.17 Understandably, this complexity makes glycomics analysis very challenging.

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Numerous techniques have been developed for interrogating the glycome at various levels 18-

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21

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environmental studies, advance spectroscopic analyses such as matrix-assisted laser

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desorption/ionization-time-of-flight mass spectrometry (MALDI-TOF/MS) as well as 1D and

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2D nuclear magnetic resonance (NMR) spectroscopy have been used to reveal important

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structural features of the exopolysaccharides in wastewater treatment systems such as aerobic

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granular sludge23,24, anaerobic granular sludge25, and membrane bioreactors (MBRs)26.

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However, MALDI-TOF/MS and NMR spectroscopy are expensive both in terms of

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investment and running costs. In contrast, high resolution gas chromatography mass

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spectrometry (GC-MS) is inexpensive, and one of the more popular tools27 as well as a

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primary technique for characterizing the structure of individual glycans28 when only small

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quantities are available, as is usually the case in environmental samples. Another great

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advantage of GC-MS glycan profiling is that multiple glycans of any given subtype can be

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characterized at once, increasing its throughput. Interestingly, although this technique is

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comparatively powerful and relatively economical, it has not been applied to analyze free

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EPS in wastewater treatment systems as far as we are aware.

, however, no single technique can define all aspects of the glycome.22 In terms of


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It is known that several bacterial EPS are homopolysaccharides, but most are

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heteropolysaccharides that consist of a mixture of neutral and charged sugar residues.29 There

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seems to be general agreement that the exopolysaccharides found in wastewater treatment

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systems contain an acidic moiety.23-25,30 In this work we provide a detailed protocol for an

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analytical procedure that can be used routinely in a reasonably equipped laboratory to

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simultaneously determine both the neutral and acidic monosaccharides, how each residue is

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linked, and the possible physical properties associated with the free EPS produced from a

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mixed culture during anaerobic digestion (AD). It also seems as if a change in nutrients can

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have a significant effect on EPS production and composition31. Until recently, research has

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mainly focused on the effect of manipulating external parameters such as nutrient levels and

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imbalances in the C:N:P ratio on the global production of EPS6,31 in a “black-box” approach.

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It is striking that fundamental research on EPS is still in its infancy, and systematic research

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on this topic is still sparse in spite of its obvious importance. Besides deducing the fine

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structure of EPS, we also provide insights into changes in glycan composition with changes

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in feed composition. Identification of the composition and characteristics of these free EPS

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could be useful to clarify various contradictions about EPS in previous studies30. After the

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composition of EPS is analyzed in detail, understanding the factors influencing its production

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would be useful to manipulate the EPS content in microbial activities, and thus improve the

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performance of anaerobic membrane reactors.

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2. Materials and Methods

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Reagents and chemicals

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Given the variety of basic carbohydrates potentially available to feed the reactors, we decided

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to use the ones commonly used in synthetic feeds such as glucose, and sucrose, which is a

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disaccharide of both glucose and fructose. In addition, to challenge the anaerobic culture we



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also chose fructose which is not commonly used, but constitutes half of the sucrose, and

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represents the other half of the carbohydrate spectrum. All analytical grade chemicals and

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biological reagents were purchased from Sigma-Aldrich, Singapore. Solvents were of GC-

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MS grade or equivalent and purchased from Merck, Singapore. Ultrapure water was obtained

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from a MilliQ water process (Millipore Advantage A10).

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Reactor operation and general parameter

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Six batch reactors with an effective volume of 5.0 L were employed for the study, and fed

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different substrates at a 7-day retention time over a period of 35 weeks to allow for “steady

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state” to occur, and their setup can be found in the Supporting Information (SI). Other

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general parameters analysed for the 6 reactors at 7-day intervals can also be found in the SI

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accompanying this paper.

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Sample preparation

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The workflow in a typical MS-based glycomics analysis comprises three stages, from sample

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preparation to data acquisition and then data analysis, preceded first by strategic planning.

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This involves a thoughtful consideration of the sample sources, and therefore what kind of

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glycans to be expected, since these ultimately will affect the sample preparation steps. The

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strategy described in this article can be found in Figure 1. Because of the work involved in

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this type of analysis it was decided that rather than take a time series of possibly changing

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samples, we would wait until “steady state” to complete one set of detailed duplicated

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analyses after 35 weeks.

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Initially, at week 35, the supernatant obtained from respective reactors was centrifuged at

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3600 g at 4 oC for 15 min, and then filtered through 10 µm filter paper (Sartorius) to remove

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sludge flocs and obtain a clear supernatant, which was then lyophilized. Two separate


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supernatant samples were taken and analysed independently to result in duplicate analyses. It

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is believed that the main components of the supernatants consist of “protein-like” compounds,

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carbohydrates, lipids, genetic materials, humic substances and small molecules.32-34 In order

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to eliminate contaminating proteinaceous materials and nucleic acids, proteinase K, DNase,

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and RNase were added prior to the extraction step.35,36; proteinase K (100 µg/mL) was added

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to the re-solubilized supernatant, and the tubes were kept at 65 oC for one hour. The mixture

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was subsequently treated with DNase (50 µg/mL) and RNase (50 µg/mL) in the presence of

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10 mM MgCl2 and 4 µL/mL chloroform, and incubated at 37

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enzymatically-digested residue was separated by centrifugation at 20000 g at 4 oC for 40 min,

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and then delipided by successive washings with methanol, methanol:chloroform (1:1, v/v)

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and chloroform. The supernatant was finally dialyzed against water (molecular weight cutoff

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7000 Da) for 1 day at 4 oC, and lyophilized before derivatization.


o

C overnight. The


148 149 150 151 152 153 154 155 156 157 158

Figure 1: An intergrated workflow for the MS-based glycomics analysis empolyed in this study (A): The specific strategies employed in every stage of sample prepartion and derivatzation. (B): Methods for the conversion of uronic acids containing polysaccharides to their corresponding PMAAs. Also shown in red is the simultaneous labelling of the reduced uronic acid with deuterim. The hydroxyl group not involved in the glycosyl-linkage are methylated with blue. After TFA hydrolysis, the partially methylated saccharides are released and reduced with NaBD4, which concurrently opens the cyclic ring to form the alditol and tag the C1 atom with a red deuterim atom. To increase the volatility of the derivatives, C atoms involved in the glycosidic linkage and the ring are acetylated (purple in colour) with acetic anhydride.

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Derivatization

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The fluffy white lyophilized sample (5 mg) was dissolved in 1 mL of ultrapure water before

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200 µL of 0.2 M 2-(n-morpholino)ethanesulfonic acid (MES) and 400 µL of 500 mg/mL


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carbodiimide reagent were added. The solution was vortexed and incubated for 3 h at 30 oC.

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Four milliliter of ice-cold 4 M imidazole-HCl and 1 mL of freshly prepared 30 mg/mL

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NaBD4 were added to the solution on ice at 5 min intervals for the first two additions; the

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third addition was performed after 2h. Excess reductant was destroyed by slowly adding 500

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µL of glacial acetic acid until the fizzing ceases, and the solution was then dialyzed against

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water overnight at 4

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monosaccharide linkage composition analysis.

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The first step of glycosylic linkage analysis is methylation; each carboxyl-reduced sample

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was dissolved in 400 µL of dimethyl sulfoxide (DMSO), and 300 µL of DMSO-NaOH

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suspension (120 mg/mL NaOH) was then added to the sample solution. After the sample was

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stirred at room temperature for 10 min, 130 µL of methyl iodide was added slowly with a

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syringe and the mixture stirred vigorously for 10 min; methylation was terminated by adding

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1 mL of water. The sample was then extracted by adding an equal volume of

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dichloromethane (DCM) to the reaction mixture; the aqueous layer was then removed and

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discarded while the organic layer was washed with 3 mL of water three times. The DCM

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phase was then dried under a stream of nitrogen.

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Next, the acid hydrolysis was performed; to each sample was added 1 mL of 2.0 M

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trifluoroacetic acid (TFA) containing 0.1 mg/mL of myo-inositol as an internal standard. The

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tubes were capped, vortexed, and incubated at 120 oC for 1 h, and then dried under nitrogen

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at 30 oC. After TFA hydrolysis, the residue was re-suspended in 10 mg/mL solution of

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NaBD4 in 2 M NH4OH (500 µL). The sample was mixed and left at room temperature 90

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min; excess NaBD4 was decomposed by the addition of 250 µL acetone, and the solvent was

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evaporated with a flow of dry nitrogen.

o

C and lyophilized; the sample can be stored frozen until



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The last step of derivatization was acetylation to form alditol acetates. Each sample was

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dissolved in 100 µL glacial acetic acid, and 500 µL ethyl acetate and 1.5 mL acetic anhydride

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were added before mixing. Acetylation was catalyzed by the addition of 58 µL perchloric

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acid (60%). After 5 min at room temperature, 5 mL of water and 100 µL 1-methylimidazole

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were added to each partially methylated alditol acetate (PMAA) sample and then mixed to

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decompose excess acetic anhydride. Once the tubes were cooled to room temperature, each

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was extracted with 2 × 1 mL DCM. The combined DCM was washed with 4 mL of water

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thrice. Following the last extraction, DCM was carefully transferred and injected into the GC-

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MS for analysis.

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Instrumentation

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A QP2010ULTRA GC-MS (Shimadzu) was used with high purity helium as a carrier gas at a

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constant flow rate of 1 mL/min; all chromatographic separations were performed on a BPX70

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column. The injector and detector temperature were maintained at 240 oC, and a split

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injection (1/10) was used with 1 µL; the column was washed five times with solvent between

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samples. The oven temperature was programmed from 140 oC, held for 2 min after sample

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injection; increased to 170 oC at 10 oC/min, held for 2 min; and then ramped to 320 oC at 6

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o

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the MS source was maintained at 230 oC. The mass spectra were recorded in the positive-ion

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electron ionization (EI) mode with acquisition range of m/z 100 to 350 at 2.14 scans per 1

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second and with a solvent delay of 3 min. The chromatographic peaks were identified using

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the NIST11 library (National Institute of Standards and Technology, Gaithersburg, MD, USA,

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http://www.nist.gov/srd/ mslist.htm).

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Preparation of standards

C/min and hold for 10 min. The total run time was less than 45 min, and the temperature of


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Seven commonly available monosaccharides (each 0.15 mol of glucose, mannose, galactose,

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xylose, arabinose, fucose and rhamnose) were weighted out into separate Teflon-lined screw-

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capped tubes. Two milliliters of 1 M methanol-HCl was added to each sample, and the tubes

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capped tightly and heated for 90 min at 80 oC. The samples were cooled to room temperature

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followed by the addition of 200 µL anhydrous 2-methyl-2-propanol to each tube; they were

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then heated to 40 oC with a stream of nitrogen to concentrate. Without pre-reduction, all the 7

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methyl glycoside samples were subjected to partial methylation separately; they then

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underwent partial acetylation, hydrolysis, reduction, acetylation, and GC-MS analysis

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identical to those described above. Relative retention time (Rf) was assigned to these partial

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methylated, partially acetylated alditol acetate standards (SI Table S2). The retention time

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(Rt) of myo-insoitol hexacetate was used as a guide to deduce the type of monosaccharide

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linkage in the actual sample, while the area of the peaks was used to quantify the relative

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molar value of the individual monosaccharides. Statistical analysis of the duplicate samples

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were performed using the Student’s t-test in Excel.

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3. Results and Discussion

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Strategic planning

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Although particulate polysaccharide determination has been improved greatly in the last

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decade, dissolved carbohydrate measurements are analytically challenging. Difficulties begin

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with quantitatively extracting and concentrating them from an aqueous sample with a high

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concentration of non-target moieties, putting pressure on the sensitivity of the analytical

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technologies. Additionally, carbohydrates exhibit multiple charge states including neutral,

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positively charged and negatively charged, making the isolation of these molecules difficult.

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No universal methodology for the rapid and reliable identification of glycan structures is

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currently available, however, a good workflow (Figure 1) will benefit from a knowledge of



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the range of glycans expected. Many bacteria produce EPS containing acidic sugars such as

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galacturonic acid and glucuronic acid.23-25,30 Although uronic acids can be detected by

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colorimetric methods such as carbazole37 or hydroxybiphenyl38,39, they often give a global

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estimate and do not differentiate between individual moieties40,41. The acidic moieties, on the

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other hand, are not detectable through acetylation and require pre-reduction to their neutral

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monosaccharide counterparts.42 Reduction of carboxyl groups usually requires activation and

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subsequent reduction; the free acidic group has to be activated first by carbodiimide and

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reduced with sodium borodeuteride (NaBD4) to generate 6,6’-dideuterio-saccharides43, which

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can be distinguished from neutral glycans by GC-MS as the presence of fragment ions with

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increased mass (M+ + 2). Besides providing a link between mass and composition, other

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advantages of using GC-MS for polysaccharide compositional analysis are its robustness and

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high resolution. Since various derivatization reagents have been reported in the literature44,

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linkage analysis is also possible if the glycan is derivatized by pre-O-methylation before

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hydrolysis.

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Methylation is an important tool for the elucidation of polysaccharide structures.45 In our

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procedure all the free hydroxyl groups of the glycans are methylated, and following TFA

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hydrolysis, the partially methylated saccharides released are further reduced and acetylated to

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yield PMAAs, which can be easily separated, identified and quantified by GC-MS. There are

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many ways to perform methylation with methyl iodide.46,47; we used a revised version of the

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Ciucanu and Kerek48-50 method as it generates cleaner chromatograms, and does not require

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the bulk generation of unstable and potentially explosive methylsulfinyl carbanion. In this

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case, NaOH-DMSO slurry, a powerful base, will deprotonate the high-pKa hydroxyl groups

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on the monosaccharide residues. Once ionized, the saccharides are treated with CH3I, which

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results in the complete methylation of the hydroxyl groups not involved in the glycoside

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linkage. Notwithstanding, methylation analysis is more qualitative than quantitative since it is


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difficult to obtain standards for each individual monosaccharide derivative. Nonetheless,

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standards are extremely important and they provide both retention time and mass spectral

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data that are necessary for the identification of derivatives. For more complex and unknown

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samples, such as in the case of environmental analysis, it is advised to generate a series of

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partially methylated, partially acetylated monosaccharide standards from their methyl-

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glycosides.

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Both quantitative and qualitative analysis of monosaccharide composition are possible by

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derivatization to alditol acetates after pre-O-methylation. The partially methylated

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saccharides will first be subjected to acid hydrolysis, which is crucial and can vary depending

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on the nature of the sample and its component glycans. Two of the most common reagents for

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hydrolysis are TFA and sulfuric acid; sulfuric acid is a harsher acid and better suited than

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TFA for complete hydrolysis. However, the presence of sulfuric acid is troublesome since the

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entire mixture is used in the subsequent steps. It can be removed, for example, by

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precipitation with barium hydroxide, but it is not practical when the sample is in micrograms.

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TFA, on the other hand, is volatile and therefore can be easily removed by evaporation. After

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acid hydrolysis the monosaccharide residues are released, but they need to be reduced with

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either NaBH451 or NaBD4 before acetylation; NaBD4 was chosen because it simultaneously

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opens the cyclic ring to form the alditol, and tags the anomeric carbon (C1) atom with

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another deuterium isotope. Glycomic analysis using isotopic labeling highlights the potential

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advantages of incorporating the mass labels into samples during derivatization, and relating

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spectral intensities to the relative quantities of the carbohydrates under investigation.

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In the final step of derivatization, acetylation not only increases the volatility of the

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derivatives for GC separation, but the C atoms in the glycosidic linkage and ring also carry

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the acetyl group (Figure 1). There are a variety of acetylation methods44; the simplest is the



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addition of acetic anhydride and catalysis with either 1-methylimidazole52 or 60% perchloric

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acid53. Typically, myo-inositol is used as the internal standard, and the acetates can be

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identified by their retention time relative to the internal standard. Finally, the PMAA

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derivatives can be injected into a GC-MS system directly.

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Polysaccharide composition

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One advantage of obtaining monosaccharide linkage composition is that it gives us the

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potential to estimate the relative proportion of different saccharides from a single analysis.

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The accuracy of this estimation depends on prior knowledge of existing classes of bacterial

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polysaccharides that are likely to be present. It will also depend, amongst others things, on

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the taxonomic origin of the bacteria and sample treatment. In the case of biological treatment

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systems, it is impossible to identify the origin of the specific microorganisms producing a

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specific carbohydrate in a mixed culture. Nevertheless, information obtained through the

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protocol discussed above can still be used to assign monosaccharide linkages, and thus

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associate them with the possible free EPS present in various reactors, as shown in Table 1.

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Although the theoretical number of structures that can be derived from combining a certain

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number of monosaccharides is high17, fortunately the actual number of structures naturally

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occurring is significantly smaller, and there are often nested structures, reflecting the limited

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number of monosaccharides and specific biosynthetic machinery present in a given biological

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context. Table 1 shows the descending ranking of various monosaccharide linkages presented

301

as a percentage, and most of the saccharide monomers are neutral glucose (Glcp), galactose

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(Galp), mannose (Manp), and rhamnose (Rhap) residues, and the polyanionic glucuronic acid

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(GlcpA) residue with various linkages.


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Table 1. Monosaccharide linkage composition with possible polysaccharide characteristics based on different feed compositions. Reactor

1

2

3

4

5

6

Major Feed Constituent

Glucose

Glucose, Urea

Glucose, NH4Cl

Fructose

Sucrose

None

C/N ratio (mg/L)

2000:0

2000:200

2000:200

2000:0

2000:0

None

Total free EPS concentration (mg/L±SD)

11.0 ± 0.35

12.0 ± 0.35

18.2 ± 0.15

9.45 ± 0.15

10.7 ± 0.25

1.4 ± 0.15

Statistically significant difference in total against reactor 1 (p = 0.05)

-

No

Yes

Yes

No

Yes

(25.6±0.3%)

(22.4±1.1%)

(28.8±0.5%)

(24.4±0.2%)

(26.3±0.1%)

(23.6±0.1%)

(24.3±0.6%)

(19.6±0.4%)

(27.6±0.4%)

(22.8±0.2%)

(24.7±0.2%)

(21.9±0.2%)

(22.4±0.6%)

(18.5±0.3%)

(13.0±0.3%)

(21.4±0.1%)

(22.8±0.3%)

(20.4±0.2%)

(9.7±0.4%)

(14.0±0.2%)

(10.5±0.3%)

(12.5±0.3%)

(8.4±0.2%)

(18.5±0.5%)

(7.9±0.1%)

(7.8±0.2%)

(6.5±0.5%)

(9.7±0.1%)

(5.8±0.1%)

(2.2±0.1%)

Monosaccharide (Mol % ± SD)

15 ACS Paragon Plus Environment


Total Monosaccharide (Mol % ± SD)

Physical properties

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89.9±0.8%

82.3±0.6%

86.4±1.0%

90.8±0.6%

88.0±0.9%

86.6±0.6%

soluble

soluble

Viscous solution with a loose gel-like behaviour

soluble

slightly viscous

soluble

Note: The number in the parenthesis refers to the average value (n = 2) and the number after the (±) symbol refers to the standard deviation (SD)

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Due to the diversity of linkage types, bacterial EPS presents a wide range of physical

306

properties. It is these linkages in polysaccharides, coupled with a variation in the monomer

307

sequence, that produce a diverse range of possible structures54. The effects which certain

308

monosaccharide residues confer on their physical properties can be seen in Table 1. It is clear

309

that these physical attributes could be determined based on the chemical composition and

310

structural niceties of the polysaccharides. For example, a composition of β1→4 or β1→3

311

linkages may confer considerable rigidity, while α1→2 linkages may yield more flexible

312

structures.29 Furthermore, it has also been suggested that the presence of a significant amount

313

of linear neutral monosaccharides with β1→4 or β1→3 linkages tends to decrease the

314

solubility of the polysaccharide, and with the presence of the right amount of uronic acid

315

residues they yield viscous aqueous solutions29,55, as in the case of reactor 3. The uronic acid

316

residues modify the original stereoregularity and helical conformations in solution; their

317

altered semi-rigid characteristics is further affirmed in the presence of several monovalent or

318

divalent cations.55 As the molar ratio of ionic substituents increases, the solubility of the

319

polysaccharide is amplified because of the strong ionic and electrostatic interactions.

320

Nevertheless, it is also not uncommon that 15-20% of the linkages are not assigned because

321

of the varied and dynamic nature of their polysaccharide structures, and experimental

322

analytical errors, and this can be seen in Table 1 where the total top five monomeric

323

structures only sum to 80-90%. It is also notable that the information generated from this type

324

of analysis is indicative only, and should not be used to assign absolute composition. Despite

325

this, it is a relatively simple and effective measure of free EPS composition that can be used

326

as the basis for further in-depth analysis.

327

Physiological aspects of EPS production



328

The formation and excretion of EPS is a process requiring considerable amounts of energy,

329

and is favored by growth under conditions of a plentiful, and readily utilizable carbon source.

330

The extent of EPS production could depend on the types of precursors, and also on other

331

physiological conditions employed. Since the biomass concentration in each reactor over the

332

35 weeks was relative constant, the amount of free EPS should not change based on biomass

333

variations. However, when there is no freely available energy source, microorganism stop

334

producing EPS, as can be seen in reactor 6 where there is no carbon source, and the free EPS

335

was very low (1.4 ± 0.15 mg/L). This presumably can be attributed to the conservation of

336

energy in preference to EPS synthesis.

337

Carbohydrates are widespread in nature and serve as initial substrates for bacteria in AD. On

338

a molecular basis, the initial steps in the EPS biosynthesis pathway essentially follow Enter-

339

Doudoroff56, and the substrates are metabolized intracellularly to intermediate compounds -

340

“sugar nucleotides” that serve as precursors, and in turn form energy-rich monosaccharide

341

donors for EPS synthesis and provide a means for interconversion.15,55 It was found that the

342

amount of total free EPS produced in the fructose-fed reactor (reactor 4) was statistically

343

lower (95% confidence level (n=2, p = 0.05) than in the glucose-grown cultures (reactor 1),

344

while the sucrose-fed (reactor 5) appeared lower, but was not statistically significant. Given

345

that sucrose is a disaccharide composed of fructose and glucose, this means during its

346

hydrolysis and metabolism half the feed is actually glucose, and hence it is perhaps not

347

surprising that the total free EPS produced from sucrose was not very different from the pure

348

glucose feed. In addition, only one of the constituent monosaccharide was statistically

349

different from those produced by glucose.

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With fructose the total free EPS was significantly different from glucose, and had a different

351

monomeric sugar composition, as observed in Table 1, and 3 out of the 5 major monomers


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produced from fructose were statistically different from glucose. It is noteworthy that these

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EPS are free and soluble, and are not cell bound; there is evidence that most of the EPS is

354

distributed on the outer layer of the cell (bound EPS), and one study even reported that the

355

EPS content on these outer layer was about four times greater than free EPS57. This further

356

suggests that the regulation of the biosynthetic pathway of EPS, or free EPS, and that of the

357

monomeric saccharide composition, both depend on the nature of the monosaccharide in the

358

feed. It was proposed that a regulatory enzyme, fructose 1,6-bisphosphtase has an activity

359

much lower than that of 6-phosphofructokinase, which is the opposite in the synthesis of

360

“sugar nucleotides”58, and this leads to a lower production of EPS in a fructose-fed medium

361

(Table 1).

362

The C source was not the only parameter affecting EPS synthesis, and it was reported that an

363

adjustment in the C/N ratio could potentially improve the characteristics of the feed and

364

produce less free EPS.12 While we did not change the C/N ratio in the feed due to the work

365

involved, our results suggest that the use of a different nitrogen feed (at a C/N ratio of 10)

366

promotes the production of more free EPS, although the addition of urea did not lead to a

367

significant increase in total free EPS (p > 0.05) in contrast to ammonia addition. It can also

368

change the monomeric sugar composition significantly, and 3 out of the 5 monomeric sugars

369

with both N sources were significantly different from glucose alone. As shown in Table 1,

370

reactors 2 and 3 had a higher estimated free EPS concentration (based on duplicates) than

371

those reactors that had no N in the feed, while the ammonium-fed reactor (reactor 3) had the

372

highest amount among all the reactors. In addition, comparison between the two N sources

373

with glucose revealed that they produced significantly different amounts of free EPS. Even

374

though the theoretical total ammonium nitrogen (TAN) was 200 mg/L for both reactors 2 and

375

3, reactor 3 had a net 200 mg/L free ammonia nitrogen (FAN). It is this FAN that diffuses

376

into the microbial cells and apparently alters the balance of intracellular activities and normal



377

material transport.31 This in turn reinforces the idea that changes in free EPS and EPS

378

composition will possibly bring about different interactions between bacteria and its

379

extracellular environment by modifying the normally negatively charged surface to repel the

380

NH4+ cations.12,31 Apparently, such action is an inherent mechanism that protects the cell

381

from fluctuations in environmental conditions.10,12,29

382

Possible implications for membrane fouling

383

EPS are one of the most important classes of membrane foulants, and may affect membrane

384

performance through pore clogging, and adhesion to the membrane surface, while leads to

385

biofouling. This enhances cake layer consolidation and results in increased membrane

386

fouling12,31 and a significant rise in energy use and overall cost.

387

The correlation between free extracellular polysaccharides and membrane fouling is

388

illustrated in Figure 2. Based on our results, it is known that most of the free EPS consist of

389

varying amounts of the negatively charged uronic acid moiety (Table 1), and monomer

390

composition changes with feed composition, and from a structural point of view, uronic acid

391

moiety is a unique component of EPS. Although present at relatively low concentrations, the

392

adsorption of free EPS will be enhanced by the ubiquitous divalent cations on the membrane

393

surface through the formation of metal-bridges with the negatively charged membrane

394

surface59,60. In addition, blocks of uronic acid moieties are able to yield an array of

395

coordination sites that assist divalent cations in their cavities, and provide a gel-forming

396

capacity (Figure 2-D). In contrast, other blocks of the free EPS that are neutral will provide

397

the chain with a flexibility and network structure during gelation. After adsorption more free

398

EPS and other extracellular polymeric substances form a gel layer which radically alters the

399

characteristics of the membrane surface61, and hence changes the membrane fouling

400

propensity. Concurrently or subsequently, free EPS will further mediate in bacterial


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attachment and reinforce the biofilm structure which is likely to be the same as that resulting

402

from EPS self-assembly62. Again, being negatively charged these organic films will

403

significantly increase bacterial attachment by hydrophobic interactions, and hydrogen

404

bonding with overlying cells63.

405 406 407 408 409 410 411 412 413 414 415 416 417 418 419 420

Figure 2. Schematic illustration of some membrane fouling mechanisms: (A) Cell biomass and EPS. The inner layer consists of tightly bound extracellular polymeric substances (TBEPS), which is consisting of free EPS and other extracellular polymeric substances that bound tightly to the cell surface. The outer layer is loose and dispersible extracellular polymeric substances, which arises from cell growth, cell lysis, or from attachment. (B) Pore clogging. Most organics, including soluble microbial products (SMPs) and colloids, could enter the membrane pores and then partially accumulate due to their “sticky nature” (i.e., characterized by adhesive force up to 8.5 nN determined by atomic force microscopy62). (C) Biofilm. Development of floc adhesion and gel layer formation emphasizing the contribution of the extracellular polymeric substances species. This will subsequently lead to the formation of a second “dynamic” membrane – cake layer. (D) Ion bridging. Extracellular polysaccharides contain an uronic acid moiety, which is negatively charged at near-neutral pHs, and will react with divalent cations such as calcium and magnesium ions to form complexes. This ion bridging mechanism can also enhance polymer entanglement, facilitating the adhesion of foulants, and strengthening the cake layer structure.

421

Impacts and prospects

422

However, no systems-level analysis of a biological process is complete without incorporating

423

the glycomics studies, and understanding how a collection of glycans are related to a

424

particular biological event such as membrane fouling. For decades most of the EPS that has

425

been isolated is from either aerobic or facultative anaerobic conditions, and there has been 21 Environment ACS Paragon Plus


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very little work on the products from strict anaerobes. Being heterogeneous in nature,

427

accurately characterizing EPS composition, and quantifying their concentration in different

428

wastewater systems is not straightforward. Their chemical composition and structure has

429

remained uncertain for a long time, and much of the knowledge of their nature has been

430

largely been based on indirect evidence.

431

Like other “omics” efforts, glycomics is being driven by new technologies for high-

432

throughput profiling. Our work not only analyzed the monosaccharide components, but also

433

finally identified their specific linkage patterns, and their modification associated with

434

changes in carbon substrates or nitrogen sources, which can have profound biological

435

consequences, is revealed. We expect our methodology will help in laying the groundwork

436

necessary for probing the structural details of EPS, and represents one step towards

437

understanding the role EPS plays in membrane fouling.

438

Nonetheless, the EPS are very complex and knowledge about them is far from complete, and

439

hence much work is still required to fully understand their precise roles in biological

440

treatment. A deeper understanding of EPS formation, and their dynamic physiochemical

441

nature, could result in new pretreatment methods for the efficient removal of EPS from the

442

supernatant, and enable novel cleaning strategies to be developed which target the membrane

443

surface. All of these new insights will ultimately improve wastewater treatment efficiency

444

and performance. Finally, the most exciting prospect of glycomic research is its potential to

445

be used in combination with prevailing genetic and biochemical tools and, when possible, for

446

integrating other data sets.

447

Acknowledgements

448

We acknowledge the financial support from the Environmental & Water Industry Programme

449

Office (PUB IDD 21100/36/6). This research grant is supported by the Singapore National 22 Environment ACS Paragon Plus

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Research Foundation under its Environmental & Water Technologies Strategic Research

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Programme and administered by the Environment & Water Industry Programme Office

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(EWI) of the PUB.

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Influence of Feed Composition on the Monomeric ... - ACS Publications

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