Effects of Microbial Degradation of Biofoulants on Microfiltration

Oct 26, 2010 - In membrane bioreactors (MBRs) for wastewater treatment, membrane fouling, particularly biofouling caused by soluble microbial products...
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Environ. Sci. Technol. 2010, 44, 8644–8648

Effects of Microbial Degradation of Biofoulants on Microfiltration Membrane Performance in a Membrane Bioreactor D A I S U K E O K A M U R A , †,‡ YOSHIHIKO MORI,† TOMOTAKA HASHIMOTO,† AND K A T S U T O S H I H O R I * ,‡,§ Department of Materials Science and Engineering and Project Research Center for Interfacial Microbiology, Nagoya Institute of Technology, Gokiso-cho, Showa-ku, Nagoya 466-8555, Japan, and Microza and Water Processing Division, Asahi Kasei Chemicals Corporation, Samejima 2-1, Fuji-city, Shizuoka 416-8501, Japan

Received July 10, 2010. Revised manuscript received October 6, 2010. Accepted October 12, 2010.

In membrane bioreactors (MBRs) for wastewater treatment, membrane fouling, particularly biofouling caused by soluble microbial products (SMP), is a nuisance problem causing decreases in permeation flux. In a previous study, we identified primary biofoulants of microfiltration (MF) membranes in SMP as polysaccharides containing uronic acids that undergo inter- and intramolecular ionic cross-linking by polyvalent cations, forming a gelatinous mass that clogs membrane pores. In the present study, we therefore attempted to isolate biofoulantdegrading microorganisms from activated sludge on a polygalacturonic acid-overlaid agar medium and evaluate their efficiency for preventing biofouling of MF membranes. Among the isolates, the fungal strain HO1 identified as Phialemonium curvatum degraded 30% of polysaccharides containing uronic acids into smaller molecules in a SMP solution containing a high concentration of saccharides after 30 days of cultivation. Microfiltration tests using a laboratory-scale submerged MBR indicated that the filtration resistance of this degraded SMP solution was lower than that of the control SMP solution without fungal inoculation. Importantly, accumulation of gelatinous mass on the membrane responsible for biofouling was avoided in the SMP solution augmented with P. curvatum HO1 during the microfiltration test. This is the first report to describe a new method for avoiding biofouling of MBRs by microbial degradation of primary biofoulants.

Introduction Membrane bioreactors (MBRs), which combine activated sludge treatment and a membrane filtration process for wastewater treatment, are gaining worldwide popularity because of their high treatment efficiencies (1, 2). However, membrane fouling is an obstacle for the efficient operation * Corresponding author phone: +81-52-735-5214; fax: +81-52735-5214; e-mail: [email protected]. † Asahi Kasei Chemicals Corporation. ‡ Department of Materials Science and Engineering, Nagoya Institute of Technology. § Project Research Center for Interfacial Microbiology, Nagoya Institute of Technology. 8644

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of MBRs, as it decreases permeation flux and therefore increases operation and maintenance costs. Membrane fouling through fiber clogging that is caused by sludge accumulation on the membrane can be prevented by mechanical methods, such as air scouring (3, 4), or through device designs (5). However, biofouling that is caused by microbial products derived from metabolism and cell lysis (6-8) is more problematic, as there are no effective measures for avoiding it. To control biofouling, biological methods, such as the control of reactor conditions including DO concentration (9) and organic loadings (10), and/or chemical methods, such as the use of cleaning agents, must therefore be employed; however, these approaches are time-consuming and costly, requiring repeated trial and error. Extracellular polymeric substances (EPS) and/or soluble microbial products (SMP) have attracted attention as causative substances for the biofouling of MBRs (11-14). EPS and SMP are complex mixtures of proteins, carbohydrates, polysaccharides, DNA, lipids, and humic substances among others, with proteins and polysaccharides in particular comprising the majority of these substances (6, 15). EPS are classified into soluble EPS, which are polymers released to bulk water, and bound EPS, which are bound to microorganisms tightly, such as capsular polymers, or loosely, such as slime (16). Laspidou and Rittmann proposed that soluble EPS are identical to SMP, although this hypothesis remains controversial (15, 17, 18). Although bound EPS can be removed together with sludge, SMP and/or soluble EPS are nuisances that are primarily responsible for biofouling (19, 20). Recently, Miura et al. suggested that members of Chloroflexi spp. are ecologically significant in MBRs used to treat municipal wastewater and are responsible for the degradation of SMP, including carbohydrates and cellular materials, which consequently reduces fouling potential of MF membranes (21). The same research group also showed that uncultured Chloroflexi spp. can degrade [14C]-labeled microbial products (22). Therefore, microorganisms that are capable of degrading the primary biofoulants in SMP could be applied for the efficient prevention of biofouling. However, there is limited information concerning the identification of the biofoulants in SMP. In the case of ultrafiltration (UF) and reverse osmosis (RO) membranes, it has been reported that proteins and polysaccharides could be the cause of biofouling (23-25). In the case of MF membranes, however, proteins are not major foulant due to their molecular sizes, which are much smaller than pore sizes (26). In a previous study, we identified the primary biofoulants of MF membranes in SMP as polysaccharides containing uronic acids that undergo inter- and intramolecular ionic cross-linking by polyvalent cations and form the gelatinous mass that clogs MF membrane pores (26). It was also reported that the presence of calcium ions decreased permeation flux through RO membranes due to the increase in the adsorption of polysaccharides (25). The aim of the present study was therefore to isolate microorganisms capable of degrading uronic acids and to evaluate their efficiency for preventing the accumulation of biofoulants on MF membranes, which are usually used in the filtration of secondary effluent in wastewater treatment.

Experimental Section Isolation of Biofoulant-Degrading Microorganisms. Two kinds of activated sludge samples from the wastewater treatment facilities of a sugar factory and a chemical factory were used as microorganism sources. For the isolation and cultivation of uronic acid-degrading microorganisms, an agar 10.1021/es102321m

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solution (1.5%) containing 1 g/L polygalacturonic acid (PGA) (powder) was overlaid onto a basal salt (BS) medium (27) including 15 g/L agar to prepare opaque PGA-BS agar plates. The activated sludge samples were diluted and then spread onto the PGA-BS agar. After cultivation at 28 °C for two weeks, microorganisms that grew on the medium were counted and expressed in terms of colony-forming units (cfu). Colonies which had formed a clear zone around their margins were selected as microorganisms that secrete uronic acid-degrading enzymes and were isolated by repeated solid culture on PGA-BS agar and liquid culture more than five times. Phylogenetic Analysis. Chromosomal DNA was extracted from selected microorganisms using a DNeasy Plant Mini Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s protocol. For the phylogenetic analysis of isolates whose colonies appeared to be fungal, the sequence of the D1-D2 region of 28S rDNA was amplified by PCR using puReTaq Ready-To-Go PCR Beads (Amersham Biosciences, Piscataway, NJ). The 28S rDNA sequence was determined using the ABI PRISM BigDye Terminator Kit (Applied Biosystems, Carlsbad, CA) and subjected to phylogenetic analysis by CLUSTAL W (28) using the neighbor-joining method (29) against the sequences of type fungal strains selected from the DDBJ/EMBL/GenBank on the basis of homology analysis by BLAST (30). The sequence datum of the 28S rDNA of the fungal strain has been submitted to the DDBJ/EMBL/GenBank under accession no. AB568605. Biofoulant Degradation Test. Artificial sewage containing 5 g/L of skim milk, 5 g/L of sucrose, 0.1 g/L of MgSO4, and 0.1 g/L of NH4Cl was continuously treated by a laboratoryscale submerged MBR (26), which was operated at a high food-microorganism ratio (F/M) of 0.2 (kg-BOD/day)/kg MLSS. When the saccharide concentration of the activated sludge in the MBR exceeded 80 mg/L, the activated sludge was filtered through filter paper with an average pore size of 1 µm (5C, Advantec Toyo Kaisha Ltd., Tokyo). The filtrate was autoclaved at 121 °C for 20 min to obtain a sterilized SMP solution containing saccharides at a concentration greater than 80 mg/L. A single colony of the uronic acid-degrading microorganism on the PGA-BS agar was inoculated into 20 mL of the sterilized SMP solution in a 100 mL-flask. The solution was then shaken at 28 °C at 115 rpm for 30 days. Weekly, an aliquot from the SMP solution was sampled and the uronic acid concentration was measured. The sample was also subjected to gel permeation chromatography (GPC) to determine the molecular weight of the chemicals contained in the SMP solution. Microfiltration Tests. A porous hollow-fiber membrane made from polyvinylidene difluoride (PVDF, Asahi Kasei Chemicals, Tokyo, Japan) was used for the microfiltration tests. The hollow fiber had inner and outer diameters of 0.6 and 1.2 mm, respectively, and the average pore diameter of the membrane was 0.1 µm. Twenty-two hollow fibers of 15 cm in length were assembled to construct a membrane module with a total membrane surface area of 0.015 m2. For the preparation of test liquids, 20 mL of the culture broth consisting of the uronic acid-degrading microorganism in the SMP solution used for the biofoulant degradation tests were inoculated into 1 L of a SMP solution containing >80 mg/L saccharides in a 5 L flask, which was then incubated at 28 °C for 30 days. An uninoculated SMP solution was also incubated under the identical conditions to serve as a control test liquid. The membrane module was submerged in 600 mL of the test liquids, which were suctioned and permeated through the membrane for filtration. The flux of filtration was set to 0.9 m/day using the flow controller of a peristaltic pump (Master Flex, Cole Parmer Co., Ltd., Vernon Hills, IL). Each set of microfiltration tests consisted of seven cycles. Each

cycle consisted of filtration for 9 min and subsequent reverse filtration for 1 min. The total test time was 70 min. Filtration resistance was calculated by the following modification of Darcy’s law according to Bian et al. (31): nRf ) TMP/µJ where Rf is the filtration resistance (m-1), n is the filtration cycle, TMP is the transmembrane pressure (Pa), µ is the permeate viscosity (Pa · s), and J is the permeate flux (m3/ (m2 · s)). Analytical Methods. Saccharide concentrations were determined by the phenol/sulfuric acid method (32). Uronic acids were analyzed using PGA as the standard, as described previously (26). Protein concentration was determined by the Bradford method using bovine serum albumin (BSA) as a standard. The molecular weight of the chemicals contained in the SMP solution was determined by GPC, as described previously (26). Total organic carbons (TOC) were quantified using a TOC analyzer (Shimadzu, TOC-V series, Japan). Fourier transform infrared (FT-IR) spectrometry analysis of a gelatinous mass was performed using Nicolet 6700/ Continumm (Thermo Scientific, West Palm Beach, FL) with a spectrum resolution of 4 cm-1.

Results and Discussion Isolation of Microorganisms Degrading Polygalacturonic Acid. To isolate microorganisms capable of degrading primary biofoulants in SMP, microorganisms were screened from activated sludge obtained from two wastewater treatment facilities on agar plates containing PGA, which is composed of only a uronic acid, as the sole carbon source. The numbers of microorganisms isolated on the PGA-BS agar were 1.1 × 105 and 1.5 × 105 cfu/mL from the activated sludge samples from the sugar factory and chemical factory treatment facilities, respectively. In this initial screening, we obtained eight colonies that formed a clear zone around their margin on the PGA-BS agar. All of these isolates were obtained from the activated sludge of the chemical factory, resembled fungal colonies, and appeared to represent two different types of fungal species. Therefore, representative isolates with these two types of colony morphologies were selected and isolated as pure strains (strains HO1 and HO2) by repeated selection on PGA-BS agar and subsequent dilution in PGA-BS liquid medium. However, although purified strain HO1 formed a clear zone around its colonies on PGA-BS agar, purified strain HO2 became incapable of forming a clear zone despite retaining the capability for growth with PGA as the sole carbon source (Figure 1). Under pure culture conditions, strain HO2 might therefore be unable to secrete a sufficient amount of PGA-degrading enzyme to form a clear zone. In contrast, strain HO1 clearly possesses the ability to stably secrete the enzyme extracellularly for the utilization of PGA. Examination of the Biofoulant Degradation Ability of the Isolates. The biofoulant degradation ability of the two isolated strains was examined using a SMP solution containing >80 mg/L of saccharides (Figure 2). The uronic acid concentration in the SMP solution gradually decreased during the cultivation of strain HO1 and was reduced to 80% of the original concentration after 30 days (Figure 2A). However, uronic acids were hardly degraded by strain HO2 during the same period of cultivation. It was also observed that the height of the GPC peak corresponding to the primary biofoulants in the SMP solution with molecular weights of 106 to 108 Da (26) decreased to 70% of the initial peak height during 30day cultivation of strain HO1 (Figure 2B). In contrast, strain HO2 showed only a 15% decrease in the height of this biofoulant peak determined by GPC during the same cultivation period. Thus, these results demonstrate that strain HO1 secretes an extracellular enzyme that degrades polysacVOL. 44, NO. 22, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 1. Colonies of two microbial strains, HO1 (A) and HO2 (B), isolated from activated sludge on PGA-BS agar plates. A clear zone can be seen around the HO1 colony (A). charides containing uronic acids, which are the primary biofoulants in SMP. Identification of Strain HO1. Morphological observation of the colonies formed by strain HO1 suggested that it was a fungal strain (Figure 1A). The filamentous hypha structure of the fungus with a thickness of about 2 µm was also confirmed by microscopy (Figure S1, Supporting Information). For conclusive identification of this isolate, the 28S rDNA D1-D2 sequence of strain HO1 was determined and phylogenetic analysis was performed. Strain HO1 was phylogenetically identified as Phialemonium curvatum, showing 100% homology of the 28S rDNA sequence in the D1-D2 region with the type strains CBS102172, CBS102173, and CBS505.82 (accession no. AB278181-278183) (Figure S2, Supporting Information). P. curvatum was previously reported to degrade lignin and pectin that is composed of polygalacturonic acids (33) and is considered to be a potential candidate for delignification in the pulping process (34). Effectiveness of Uronic Acid Degradation for Preventing Biofouling. We next investigated the effectiveness of the degradation of uronic acids by P. curvatum HO1 for preventing biofouling of an actual MBR. The test liquids were prepared by a 30-day incubation of the SMP solution containing a saccharide concentration of more than 80 mg/ L, which typically causes the accumulation of a thick gelatinous mass on MBR membranes during microfiltration tests (26), with and without (control) inoculation with a 1/50 volume of P. curvatum HO1 culture solution. As shown in Table 1, the concentrations of saccharides and uronic acids were 73 and 21 mg/L, respectively, in the presence of P. curvatum HO1 and were significantly lower than the con8646

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FIGURE 2. Time profiles of uronic acid concentrations (A) and GPC peak height of the primary biofoulants (B) in the SMP solution during cultivation of strains HO1 and HO2 for 30 days at 28 °C. C, uronic acid concentration during incubation; C0, original uronic acid concentration; H, foulant GPC peak height during incubation; H0, original foulant GPC peak height. The experimental results are the means (symbols) ( standard errors (error bars) obtained from independent measurement of three samples.

TABLE 1. Effects of Degradation of Biofoulants on Microfiltration

TOC (mg/L) pH (-) saccharides before filtration (mg/L) after filtration (mg/L) gelatinous mass (mg/L) uronic acids before filtration (mg/L) after filtration (mg/L) gelatinous mass (mg/L) filtration resistance ×10-11 (m-1)

control, SMP solution

degraded, SMP solution

89 ( 0.45 6.8

82 ( 0.40 6.9

105 ( 0.50 49 ( 0.25 2700 ( 20

73 ( 0.35 17 ( 0.23 -

31 ( 0.15 8.9 ( 0.10 520 ( 10 5.6

21 ( 0.20 8.9 ( 0.10 3.4

centrations in the absence of P. curvatum HO1 (105 and 31 mg/L, respectively). This implies that strain HO1 degraded 30% of the saccharides and uronic acids in the SMP solution during the 30-day incubation. GPC analysis of these test liquids also revealed that the peak area for polysaccharide biofoulants at molecular weights of 106-108 Da in the SMP solution with P. curvatum HO1 decreased by 30% compared with the peak in the control SMP solution (Figure 3). In addition, the peak at molecular weights of 103-104 Da, which is considered to correspond to degradation products of polysaccharides, intensified 3-fold in the SMP solution containing P. curvatum HO1. Thus, the results obtained from the examination of the biofoulant degradation ability of the isolates and those of the test liquid preparations for microfiltration tests were reproduced, even though the experi-

FIGURE 3. GPC chromatogram of the microfiltration test liquids prepared by incubation of a SMP solution in the presence of P. curvatum HO1 (degraded SMP solution) or in the absence of the fungal strain (control SMP solution). mental conditions such as the culture volume and inoculation method differed. These results indicate that PGA-degrading P. curvatum HO1 can partially degrade polysaccharides containing uronic acids into smaller molecules in the SMP solution obtained from MBR operation by extracellular enzymes. The test liquid resulting from the partial degradation of polysaccharides by P. curvatum HO1 was referred to as degraded SMP solution and was further subjected to a microfiltration test, as was the control SMP solution without microbial degradation. TOC concentrations of the control and degraded SMP solutions were 89 and 82 mg/L, respectively, while their saccharide portions in TOC were calculated from saccharide concentrations as about 42 and 29 mg/L, respectively. The less decrease in TOC than in saccharide concentrations suggests that a small amount of soluble organic compounds other than saccharides were produced by cultivation of P. curvatum HO1, such as proteins and DNA. At least, enzymes that are responsible for polysaccharide degradation are thought be secreted by this degrader, as described above. During the microfiltration test of the degraded and control SMP solutions, which consisted of seven cycles of filtration and backwash of the test liquids, changes in the TMP were monitored (Figure 4). The TMP in the MBR with the degraded SMP solution was >10 kPa lower than that with the control SMP solution. As a result, the filtration resistance values of the tests with the degraded and the control SMP solutions were 3.4 × 10-11 and 5.6 × 10-11 m-1, respectively (Table 1). To confirm that the reduction of the filtration resistance in the degraded SMP solution was significant, directly reflecting the biofoulant concentration decreased by microbial degradation, the control SMP solution was diluted with distilled water and subjected to the microfiltration test (Table S1, Figure S3, Supporting Information). As a result, a linear correlation was confirmed between filtration resistances and saccharide concentrations obtained from the control, degraded, and diluted SMP solutions (Figure S4, Supporting Information), as shown previously (26). Thus, it was clearly demonstrated that the addition of P. curvatum HO1 to the test solution can reduce filtration resistance through the degradation of biofoulants. Importantly, no visible residue was observed on the membrane after the microfiltration test of the degraded SMP solution, whereas a thick gelatinous mass accumulated on the membrane surface in the case of the control SMP solution. The saccharide and uronic acid concentrations of this gelatinous mass were approximately 25- and 17-fold higher, respectively, than those of the original test liquid of the control SMP solution (Table 1). Such gelatinous masses are formed by the cross-linking of single polysaccharide chains with polyvalent cations at uronic acid units, which clog membrane pores resulting in biofouling (26). However, FTIR spectrometry analysis of the gelatinous

FIGURE 4. Effect of biofoulant degradation on transmembrane pressure (TMP) during microfiltration tests. (A) Control SMP solution; (B) degraded SMP solution by P. curvatum HO1.

mass showed peaks probably derived from proteins or aminosugar units as well as polysaccharides (Figure S5, Supporting Information). In fact, 50 mg/L of proteins were contained in the gelatinous mass as minor components. On RO membranes, alginate also forms a crossed-linked gel layer with calcium ions and causes biofouling (35), and adsorption of polysaccharides onto RO membranes is greatly enhanced by calcium ions (24, 25). In addition, there is initial synergistic fouling effect when RO membranes are fouled by both BSA and alginate, as compared to fouling by BSA or alginate alone (24). Therefore, proteins contained in wastewater might also adsorb to the gelatinous mass when gel was formed by crosslinking of polysaccharides containing uronic acids. The similarity in the fouling mechanism between MF and RO membranes suggests that our approach, microbial degradation of uronic acids, might also be effective for preventing biofouling not only of MF membranes but also of RO membranes. In summary, the accumulation of gelatinous masses responsible for biofouling can be prevented by the PGAdegrading microorganism P. curvatum HO1, even if the degradation of polysaccharide is only partial. This is the first report to describe a new method for avoiding biofouling of MBRs by microbial degradation of primary biofoulants. We are now attempting to isolate microorganisms that are capable of degrading biofoulants more efficiently and rapidly than P. curvatum HO1. We expect that cultivation of these microorganisms in an appropriate nutrient-rich medium at the outside of a MBR enables the augmentation of enough microbial cells into the MBR for rapid degradation of biofoulants, resulting in effective prevention of membrane fouling.

Supporting Information Available Additional information as noted in the text. This material is available free of charge via the Internet at http://pubs.acs.org.

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