The importance of laboratory water quality for studying initial bacterial adhesion during NF filtration processes
Introduction
Nanofiltration (NF) and reverse osmosis (RO) membranes are commonly used for the removal of organic matter and trace contaminants, such as pesticides, during water treatment processes (Cyna et al., 2002). The efficiency of NF and RO processes is however adversely affected by membrane biofouling (Flemming, 1997; Ivnitsky et al., 2007), principally due to the formation of biofilms (Flemming, 2002). These ecosystems are usually made up of a community of dead and living microorganisms held together by a matrix of polysaccharides, lipids, proteins, organic matter, amongst other components (Flemming, 2002). Biofilms are ubiquitous in NF and RO membrane plants (Houari et al., 2009; Vrouwenvelder et al., 1998, 2008; Khan et al., 2013) and are the Achilles heel of NF and RO processes (Flemming et al., 1997) as they are difficult to remove (Hijnen et al., 2012). Biofouling increases pressure drop along the membrane module (Vrouwenvelder et al., 2009a; Hijnen et al., 2009), leading to increased costs associated with energy consumption. The presence of biofilms on the membrane surface has also been shown to significantly affect permeate flux, and solute retention (Ivnitsky et al., 2005; Huertas et al., 2008). The decrease in solute retention and permeate flux has been attributed to enhanced concentration polarisation caused by the biofilms (Herzberg and Elimelech, 2007). It has been shown that the concentration polarization also maintains the presence of biofilms by concentrating nutrients from the bulk environment (Chong et al., 2008; Vrouwenvelder et al., 2009b).
Biofilm formation is initiated by the irreversible adhesion of bacterial cells onto the membrane's surface, which is influenced by a number of factors. Firstly, the cell properties such as hydrophobicity (Ridgway et al., 1985) and cell surface charge (Subramani and Hoek, 2008) have been found to affect adhesion. Secondly, the membrane physicochemical properties (roughness, charge and hydrophobicity) have been shown to impact the degree of adhesion. In general, the rougher and the more hydrophobic the membrane is, the more cells will adhere to the surface (Subramani and Hoek, 2008; Myint et al., 2010; Khan et al., 2011). Finally, the presence of a conditioning layer on the membrane also affects bacterial adhesion (Subramani et al., 2009). A recent study has shown that a conditioning layer of salts and organic carbon promoted a homogeneous biofilm, whilst the absence of a conditioning layer resulted in a scattered and thin biofilm (Baek et al., 2011).
The intractable nature of the biofouling problem has led to a significant increase in research in this area in recent years (Herzberg and Elimelech, 2007; Chong et al., 2008; Subramani and Hoek, 2008; Baek et al., 2011; Fonseca et al., 2007). These studies range from the effects of biofilms on process performance (Ivnitsky et al., 2005; Huertas et al., 2008) to biofouling control through the design of antifouling membranes (Miller et al., 2012; Bernstein et al., 2011). Although membrane biofouling research methodologies differ from one research laboratory to another, they generally share a common pre-treatment procedure involving the compaction of the studied membrane prior to biofouling experiments. To accurately monitor flux changes and solute retention during NF and RO experiments caused by osmotic pressure or membrane fouling, membranes are purposely compacted to prevent changes due to the effect of pressure during the experiment. The compaction of NF and RO membranes is carried out under different filtration conditions depending on the laboratory they are carried out. The compaction is typically undertaken at a pressure between 6 and 25 bar and up to 18 h in duration (Herzberg and Elimelech, 2007; Baek et al., 2011; Fonseca et al., 2007; Suwarno et al., 2012). This translates into a typical water permeation volume between 2 L (membrane flux = 50 L h−1 m−2, time = 18 h and 22.44 cm2 membrane area) and 15 L (membrane flux = 65 L h−1 m−2, time = 12 h and 186 cm2 of membrane area) (Baek et al., 2011; Suwarno et al., 2012) calculated as: V (L) = Flux (L/h m2) × time (h) × Membrane area (m2).
Although membrane compaction is a prerequisite to most NF and RO experimental studies, including bioadhesion/biofouling, the type of water used to compact the membrane may vary considerably from one laboratory to another. The water used in recent published studies on initial adhesion and biofouling experiments spans from non-sterilised tap water (Hijnen et al., 2009; Khan et al., 2011; Vrouwenvelder et al., 2009c, 2007; Botton et al., 2012; Khan et al., 2010), DI water (Huertas et al., 2008; Herzberg and Elimelech, 2007; Myint et al., 2010; Baek et al., 2011; Lee et al., 2010) and MilliQ water (Chong et al., 2008, 2007; Pang et al., 2005). Tap water and DI water will vary in quality depending on the water source and the yearly season (Gibbs et al., 1993). In essence, the total carbon, biological and endotoxin contents will differ from one water type to another, whether the water is sterilized or not. Moreover, when considering filtration aspects, all insoluble water constituents will most certainly be deposited on the membrane surface during the compaction, thus altering the membrane surface from its original state. The conditioning layer formed during the compaction pre-treatment of NF/RO is likely to result in altered surface characteristics thereby affecting subsequent biofouling experiments.
The objective of this study was to first demonstrate the impact of the choice of water used during compaction of NF membranes in terms of membrane performance, surface characterisation and secondly, to investigate whether the water used during membrane compaction also affects bioadhesion outcomes.
Section snippets
Water source and characterisation
Three different water grades were used in this study: tap water provided by south Dublin water municipality, deionized water obtained by a purifying water system (Elgastat B124, Veolia, Ireland) and Grade 1 pure water (18.2 MΩ cm−1) obtained from an Elga Process Water System (Biopure 15 and Purelab flex 2, Veolia, Ireland), hereafter referred to as MilliQ water. Conductivity and pH measurements were performed on all samples at room temperature (20 °C) and total organic carbon of all water
Water quality assessment
The different water qualities used in this study for compacting NF 270 membranes were characterized prior to compaction experiments and are presented in Table 1.
No detectable solids were measured in MilliQ and deionized water samples used in this study. MilliQ water had the lowest pH, total organic carbon, and conductivity values compared to deionized and tap water, respectively. MilliQ water has a very low conductivity of 0.4 μS cm−1 followed by DI water with a conductivity of 4 μS cm−1. The
Discussion
The aim of this study was to investigate the effects of laboratory water quality during compaction of nanofiltration membranes in terms of performance, surface property changes as well as its influence on standard bio-adhesion assays.
Filtration performance together with the physicochemical, physical properties and biological assessment of NF 270 membranes were analysed following 0.5 L, 2 L and 5 L set permeation volumes during compaction with different water sources. Dynamic bioadhesion assays
Conclusion
The impact of laboratory water quality was assessed following compaction of the NF 270 membrane by analysing the membrane performance and surface characteristics, as well as the adhesion characteristics of P. fluorescens. Tap and DI water compaction resulted in a cake layer on the membrane surface consisting of living and dead bacteria and diatoms, organic matter, dissolved solids and other components, as these were present in the water used for compaction. There was a clear difference in the
Acknowledgements
This research was supported by the European Research Council (ERC), project 278530, funded under the EU Framework Programme 7 and also with the financial support of Science Foundation Ireland under Grant Number “SFI 11/RFP.1/ENM/3145. The authors would like to thank Dr. Dennis Dowling and the Surface Engineering research group at UCD. We thank Dr. Ian Reid of the NIMAC microscopy platform UCD. We thank Mr. Pat O'Halloran for his invaluable technical assistance, and Mr. Liam Morris for the
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Both authors contributed equally to this work.