Highly Efficient Enzyme-Functionalized Porous Zirconia Microtubes for

Article pubs.acs.org/est

Highly Efficient Enzyme-Functionalized Porous Zirconia Microtubes for Bacteria Filtration Stephen Kroll,* Christoph Brandes, Julia Wehling, Laura Treccani, Georg Grathwohl, and Kurosch Rezwan Advanced Ceramics, University of Bremen, Am Biologischen Garten 2, 28359 Bremen, Germany ABSTRACT: In contrast to polymer membranes, ceramic membranes offer considerable advantages for safe drinking water provision due to their excellent chemical, thermal, and mechanical endurance. In this study, porous ceramic microtubes made of yttria stabilized zirconia (YSZ) are presented, which are conditioned for bacteria filtration by immobilizing lysozyme as an antibacterial enzyme. In accordance with determined membrane pore sizes of the nonfunctionalized microtube of ≤200 nm, log reduction values (LRV) of nearly 3 (i.e., bacterial retention of 99.9%) were obtained for bacterial retention studies using gram-positive model bacterium Micrococcus luteus. Immobilization studies of lysozyme on the membrane surface reveal an up to six times higher lysozyme loading for the covalent immobilization route as compared to unspecific immobilization. Antibacterial activity of lysozyme-functionalized microtubes was assessed by qualitative agar plate test using Micrococcus luteus as substrate showing that both the unspecific and the covalent lysozyme immobilization enhance the microtubes’ antibacterial properties. Quantification of the enzyme activity at flow conditions by photometric assays reveals that the enzyme activities of lysozyme-functionalized microtubes depend strongly on applied flow rates. Intracapillary feeding of bacteria solution and higher flow rates lead to reduced enzyme activities. In consideration of different applied flow rates in the range of 0.2−0.5 mL/min, the total lysozyme activity increases by a factor of 2 for the covalent immobilization route as compared to the unspecific binding. Lysozyme leaching experiments at flow conditions for 1 h show a significant higher amount of washed-out lysozyme (factor 1.7−3.4) for the unspecific immobilization route when compared to the covalent route where the initial level of antibacterial effectiveness could be achieved by reimmobilization with lysozyme. The presented platform is highly promising for sustainable bacteria filtration.

(1). INTRODUCTION The access to clean water is one of the fundamental requirements for life. Today, 884 million people lack access to safe drinking water, 2.6 billion people have little or no sanitation, and millions of people die annually from diseases transmitted by polluted water.1−6 Basically, natural water can be polluted by chemical (e.g., organic or inorganic species), physical (e.g., color), and biological (e.g., bacteria, viruses) contamination. Particularly, pathogenic bacteria (e.g., Vibrio cholerae, Legionella pneumophilia, Salmonella typhi) are responsible for waterborne diseases and present a direct risk to human health.7−11 Safe drinking water is commonly provided by disinfection, for example by chlorine, ozone, and UV treatment, which has a high effectiveness in killing bacteria. However, toxic or carcinogenic disinfection by-products (DBPs) are formed, which have to be removed subsequently from the purified water samples. The utilization of membranes can overcome this problem by the retention of the bacteria cells at the filtering layer and thus providing a permeate free from bacterial contaminants and DBPs. The main advantage of membrane technology is the fact that it works without the addition of chemicals, with a relatively low energy use and straightforward process handling.12−15 © 2012 American Chemical Society

However, membrane fouling is the most critical problem in many membrane technology applications where fouling is often mediated by biomolecules (e.g., proteins) or bioorganisms (e.g., fungi, bacteria, viruses). Fouling leads to a decline of permeate flux making more frequent cleaning and replacement necessary, which then increases operating costs.16−19 Today, membranes for bacteria filtration providing a sterile barrier with a cutoff of 200 nm are widely used for process water and wastewater treatments in biotechnology, food technology, and pharmaceutical technology. To improve the antifouling behavior, antibacterial components that either kill or prevent the growth of bacteria can be immobilized onto membrane surfaces leading to self-cleaning matrices. For example, antibacterial substances are silver-based agents (silver ions and silver nanoparticles, respectively), antimicrobial peptides (AMPs such as mellitin, cecropin A), proteins (e.g., lactoferrin), and enzymes (bacterial cell wall hydrolases such as lysozyme).20−24 Received: Revised: Accepted: Published: 8739

February 16, 2012 July 11, 2012 July 24, 2012 July 24, 2012 dx.doi.org/10.1021/es3006496 | Environ. Sci. Technol. 2012, 46, 8739−8747

Environmental Science & Technology

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Figure 1. Covalent functionalization strategy for lysozyme immobilization based on membrane activation by hydroxylation (step 1), followed by silanization with APTES (step 2), and finally lysozyme immobilization (step 3).

The enzyme lysozyme (E.C. 3.2.1.17, peptidoglycan Nacetylmuramoylhydrolase) is one of the best studied and abundant antibacterial factors and its immobilization behavior to different matrices (e.g., oxide and calcium phosphate particles, polymer and ceramic membranes, ceramic micropatterns, wool, cotton/cellulose materials) is well characterized.25−31 Lysozyme has a molecular weight of 14.7 kDa, the isoelectric point (IEP) of this enzyme is 11, and one molecule has dimensions of around 3 × 3 × 4.5 nm3. Because of enzymatic activity, lysozyme can degrade the bacteria cell wall by catalyzing the hydrolysis of β-(1−4) linkages between Nacetylmuramic acid and N-acetyl-D-glucosamine residues in peptidoglycan.32,33 Compared to polymeric membranes, ceramic membranes are particularly attractive due to their excellent chemical, thermal, and mechanical stability34−36 but require generally a more sophisticated fabrication process. The objective of this work is to prepare self-cleaning zirconia microtubes for bacteria filtration by immobilizing the antibacterial enzyme lysozyme. A covalent functionalization strategy based on membrane activation by hydroxylation, followed by silanization using 3aminopropyltriethoxysilane (APTES), and finally lysozyme immobilization is developed and their performance is compared at different flow conditions.

with an electrical resistance of 18 MΩ (Synergy, Millipore, Germany) was used for all experiments. (2.2). Preparation and Characterization of YSZ Microtubes. YSZ microtubes were fabricated by extrusion as shown in our previous study27 and a sintering temperature of 1050 °C with a dwell time of 2 h was found to provide ideal membrane properties being suitable for bacteria filtration. For the determination of membrane properties, YSZ microtubes were characterized by Hg intrusion porosimetry (pore size distribution, open porosity), BET analysis (specific surface area), three-point bending test (flexural strength), and water permeate flux measurement. (2.3). Bacterial Retention Test. The bacterial retention test was performed using YSZ microtubes sintered at 1050 °C and gram-positive bacterium M. luteus as substrate (DSM Cat. No. 20030, Deutsche Sammlung von Mikroorganismen and Zellkulturen GmbH, DSZM, Germany). For the determination of bacterial retention, one individual microtube with an accessible length of 5 cm was connected with one end to a convenient silicon tubing. The other end of the tube was closed with glue (2-component epoxy resin adhesive, UHU GmbH) to ensure operating in dead-end mode. A constant flow rate of 1 mL/min for intracapillary feeding of bacteria solution was performed using a peristaltic pump (BVB Standard, Ismatec). For this purpose, M. luteus was grown from a frozen culture on BHI agar plates at RT. A bacteria inoculum from the agar plates was transferred into shaking flasks filled with 30 mL of BHI medium (37 g/L). After inoculation, the culture was grown for 24 h at RT and 120 rpm (aerobic conditions). Subsequently, the bacteria were harvested by centrifugation at 4000 rpm and RT for 10 min. The cell pellet collected by centrifugation was resuspended in 0.1 M phosphate buffer (pH 7) to a final optical density at 450 nm (OD450 nm) of 0.75 to serve as a diluted bacteria feed solution. The applied filtration time was 1 h and bacterial retention tests were done in triplicate determination based on three individual microtubes and separate bacterial cultures. Three different methods for bacteria quantification were applied to determine the ratio of the bacterial concentration between permeated volumes and feed solutions. First, the samples were analyzed by photometric measurement (VIS spectrophotometer XION 500, HACH LANGE GmbH, Germany) to determine OD450 nm. Second, a microbial cell viability assay based on luminescent adenosine triphosphate (ATP) measurement was performed to determine the number of viable bacteria cells (BacTiter-Glo assay). Therefore, a volume of 100 μL of feed and permeate solution respectively was mixed with 100 μL of BacTiter-Glo reagent and luminescence signals were measured using a platereader (Plate CHAMELEON V, Hidex, Germany). Third, the

(2). MATERIALS AND METHODS (2.1). Materials. The zirconia powder and reagents were obtained from commercial sources and used without further purification: Yttria (3 mol %) stabilized zirconia (TZ-3Y-E, Lot. Z309470P, specific surface area = 15.1 m2/g) was obtained from Tosoh, Japan. Bees wax (bleached, product number 14367, Lot. 40208217), stearic acid (95%, product number 175366, Lot. U03863-042), decane (≥99%, product number D901, Lot. MKBH7223 V), sulfuric acid (95−97%, product number 30743, Lot. SZBA0810), hydrogen peroxide (≥35%, product number 95299, Lot. 1406128), 3-aminopropyltriethoxy silane (APTES, 99%, product number 440140, Lot. 92196EJ399), acid orange II sodium salt (product number 75370, Lot. WE304250/1), lysozyme from chicken egg white (lyophilized powder ∼95%, MW 14.7 kDa, product number L6876, Lot. 028K0062), N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC, commercial grade, product number E7750, Lot. 1427110), brain heart infusion (BHI) broth (product number 53286, Lot. BCBC7611), and agar (product number A7002, Lot. 050M0202 V) were purchased from Sigma-Aldrich Chemie GmbH, Germany. Bradford protein assay (product number 500-0006, Lot. 109640) was purchased from Bio-Rad Laboratories, Germany. BacTiter-Glo microbial cell viability assay (product number G8231, Lot. 014452) was obtained from Promega, Germany. Double deionized water 8740

dx.doi.org/10.1021/es3006496 | Environ. Sci. Technol. 2012, 46, 8739−8747


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solution (2 vol %) for 15 min (amino-activated 1), 90 min (amino-activated 2), and 24 h (amino-activated 3) respectively at 75 °C under continuous shaking in a water bath (Memmert, Germany). Deionized water was chosen as solvent and the aqueous APTES solution at pH 3.5 was used. After silanization, the microtubes were removed from APTES solution and washed five times with 10 mL deionized water. Quantification of bound APTES molecules was performed by photometric acid orange II assay as described previously (duplicate determination).27,39,40 (2.4.3). Immobilization of Lysozyme. For lysozyme immobilization experiments, lysozyme was dissolved in deionized water to a final concentration of 1 mg/mL. In relation to the covalent functionalization strategy, APTESfunctionalized microtubes (amino-activated 2) were treated with freshly prepared lysozyme solution containing 0.1 M EDC for activation of carboxylic acid residues of the enzyme. The unspecific binding was performed by incubation of the untreated microtube with 1 mg/mL lysozyme solution without activation by EDC. All lysozyme immobilization experiments were carried out using 293 mg microtubes (amino-activated 2 and untreated, respectively), which were incubated with 15 mL lysozyme solution for 1 h at RT and 1000 rpm (thermomixer). After lysozyme immobilization, the microtubes were washed 5 times with 50 mL deionized water for 5 min at RT and 1000 rpm to remove unbound and weakly bound lysozyme molecules. Considering the amount of lysozyme in the wash fractions, wash out effects were analyzed. To determine the amount of immobilized lysozyme in consideration of wash out effects, a Bradford assay for protein quantification was used with lysozyme as the standard (duplicate determination).41 (2.5). Bacterial Adsorption on Functionalized Microtube Surfaces. Bacterial adsorption tests were carried out under sterile conditions using single microtubes of 10 cm (293 mg) with different functionalization status derived from the treatment with Piranha solution (hydroxyl-activated) and silanization with APTES (amino-activated type 2). Compared to an untreated microtube (reference), hydroxyl- and aminoactivated microtubes were intracapillary fed with a suspension of M. luteus (OD450 nm of 0.75) using a peristaltic pump. The applied flow rate was 0.2 mL/min and bacterial adsorption tests were performed for 10 min, 1 h, and 4 h, respectively. After passing the intracapillary space of the microtubes (lumen of 55.4 mm3), the obtained solutions compared to corresponding feed solutions were analyzed by photometric measurement (OD450 nm). From the ratio between bacteria found in the supernatant and the feed solution, the amount of adsorbed bacteria was determined as a function of adsorption time. Bacterial adsorption tests for the untreated, hydroxyl-, and amino-activated microtubes were done in triplicate. (2.6). Lysozyme Activity. The antibacterial activity of lysozyme-functionalized microtubes was qualitatively investigated by agar plate tests and quantitatively analyzed by photometric measurements using a gram-positive bacterium in the form of M. luteus as substrate. (2.6.1). Agar Plate Test. Agar plate tests were carried out using lysozyme-functionalized microtubes of 2.5 cm (73.25 mg) derived from the covalent (based on amino-activated type 2) and unspecific immobilization, respectively. An untreated microtube (no lysozyme immobilization) served as reference. Each microtube sample was separately treated with 10 mL bacteria solution (M. luteus solution diluted with sterile 0.1 M

permeated and feed solutions were analyzed by manual counting of bacteria colony forming units (CFUs) on agar plates. (2.4). Immobilization of Antibacterial Lysozyme onto YSZ Microtubes. For providing lysozyme-functionalized zirconia microtubes, a covalent functionalization strategy was developed. In the first reaction step, YSZ microtubes sintered at 1050 °C were activated by hydroxylation, and different hydroxylation methods in the form of an acidic as well as alkaline hydroxylation and a hydrothermal treatment (steam atmosphere) respectively were tested. The hydroxylated membrane was then functionalized with APTES to generate terminal amino groups on the membrane surface (reaction step 2). Finally, lysozyme molecules were covalently bound to the amino-activated membrane (reaction step 3) by additional activation of the carboxylic acid containing residues of the enzyme (aspartic acid, glutamic acid, carboxy terminus) using N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC). In summary, Figure 1 indicates the reaction steps in relation to membrane activation by hydroxylation (1), followed by silanization using APTES (2), and finally lysozyme immobilization (3). (2.4.1). Membrane Activation by Hydroxylation. For each hydroxylation method 2.5 g of sintered YSZ microtubes were used. Acidic hydroxylation was carried out using freshly prepared Piranha solution (97% H2SO4: 35% H2O2, 3:1, v/ v). Alkaline hydroxylation was based on the utilization of 15 M NaOH. Each set of YSZ microtubes was placed in individual vials and the sample tubes were placed in a thermomixer (Eppendorf, Germany) and incubated at different reaction conditions. For each hydroxylation method, the initial batches of 2.5 g of the sintered microtubes were separated into 10 samples of 0.25 g, which were individually treated with 2 mL Piranha and NaOH solution, respectively. Solutions were held at temperatures of either RT or 95 °C respectively, and the reaction time was 30 min under continuous shaking at 500 rpm. Hydrothermal treatment was carried out by incubation of YSZ microtubes in a steam atmosphere (20 min at 121 °C under pressure of 2 bar) using an autoclave (Systec GmbH, Germany). After each activation process, the hydroxylated microtubes were thoroughly rinsed with deionized water until the effluents were pH neutral and dried.37,38 To quantify the amount of hydroxyl groups on the membrane surface, potentiometric titration was performed (automatic titration unit with pH meter, DT1200, Dispersion Technology Inc., USA). Both blank sample (untreated microtube) and test samples (hydroxyl-activated microtubes) were pestled and in each case 2.5 g of crushed-membrane parts were used to perform potentiometric measurements. The membrane sample was dissolved in 35 mL deionized water and 150 μL HCl (1 M) and then the solution was titrated by standard KOH solution (0.5 M). A titration measurement was also performed without the addition of membrane material to serve as reference. Considering the difference of the titration curves (reference versus membrane-based samples) and the specific surface area of the corresponding microtubes hydroxyl group surface densities (OH/nm2) were calculated. (2.4.2). Preparation of Amino-Activated Microtubes by Silanization Using APTES. Silanization using APTES was carried out using hydroxyl-activated microtubes treated with Piranha solution at RT for 30 min (acidic hydroxylation). Each set of hydroxyl-activated membranes (10 cm = 293 mg) were incubated with 15 mL of freshly prepared 85.5 mM APTES 8741



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Figure 2. Possible setup of a capillary membrane module for operation in dead-end mode (A), single sintered YSZ microtubes fabricated by extrusion, insert: a cross section of a microtube (B) and a SEM micrograph of the obtained microstructure (C). ICS = intracapillary space.

phosphate buffer to a final OD450 nm of 0.75) for 30 min at RT under continuous shaking at 120 rpm. After this, the microtubes were directly transferred to fresh BHI-agar plates followed by incubation for 4 d at RT. Finally, bacteria growth was qualitatively analyzed by taking photographs of the agar plates. (2.6.2). Quantitative Analysis of Lysozyme Activity at Flow Conditions. The determination of lysozyme activity was based on the lysis of M. luteus cells monitoring the decrease in optical density at 450 nm (OD450 nm). To quantify the enzyme activities of lysozyme-functionalized microtubes at flow conditions, single microtubes of 10 cm (293 mg) derived from the covalent (based on amino-activated type 2) and unspecific immobilization were used, respectively. A suspension of M. luteus (OD450 nm of 0.75) was intracapillary fed to single lysozyme-functionalized microtubes (1.38 mm outer diameter and 0.84 mm inner diameter) using a peristaltic pump. After passing the intracapillary space of the microtube (55.4 mm3), the decrease in absorbance of the bacteria solution was measured photometrically at 450 nm for 10 min (time interval of 5 s) using a flow through cell (Hellma Analytics, Germany). Finally, the bacteria solution was collected in a waste reservoir. Lysozyme activities were determined at different flow rates (0.2, 0.3, 0.4, and 0.5 mL/min, respectively) based on duplicate measurements. One lysozyme unit is defined as the amount of enzyme required to produce a change in the absorbance at 450 nm of 0.001 units per minute at pH 7.0 (0.1 M phosphate buffer) and 25 °C using a suspension of M. luteus as substrate (initial OD450 nm of 0.75).42 (2.6.3). Lysozyme Leaching at Flow Conditions. Lysozyme leaching at flow conditions was analyzed using lysozymefunctionalized microtubes derived from the unspecific and covalent immobilization, respectively. According to the lysozyme activity tests, leaching experiments were performed by intracapillary feeding with a buffer solution (0.1 M phosphate buffer, pH 7) at 25 °C using a peristaltic pump. The applied flow rates were 0.2 and 0.5 mL/min respectively,

and leaching experiments were performed for 1 h. After passing the intracapillary space of the microtubes sample fractions were collected every 10 min and analyzed by Bradford assay with lysozyme as standard.41 Lysozyme leaching experiments for the unspecific as well as the covalent immobilization route were done in triplicate.

(3). RESULTS AND DISCUSSION (3.1). Properties of Extruded YSZ Microtubes. In this work, all experiments were performed with single YSZ microtubes without integrating them into modules. Figure 2 presents a possible build-up of a capillary membrane filtration module for operating in dead-end mode (A) followed by single microtubes derived from extrusion (B) and finally the resulting microstructure of the sintered microtubes (C). As described in our previous study,27 YSZ microtubes sintered at 1050 °C with a reliable cutoff of 200 nm, a high open porosity (>50%), and a sufficient mechanical stability showed promising membrane properties for bacteria filtration. This is proved by bacterial retention studies operating in deadend mode where the ratio of bacteria in the permeate and feed samples was quantified by three different methods by determining the optical density (OD450 nm), using an ATP assay and performing a CFU test. In accordance with determined membrane pore sizes, our bacterial retention studies using M. luteus result in a bacterial retention of 99.9% (OD450 nm), 99.3% (ATP assay) and 99.6% (CFU method) respectively, that is, log reduction values (LRV) of nearly 3 were obtained (Figure 3). Because of the requirements of U.S. Environmental Protection Agency (EPA) for safe and clean drinking water provision, the bacteria filter criterion of at least 6 LRV level is not achieved. Nevertheless, tailored immobilization of antibacterial components (e.g., lysozyme) to the microtube surface can help to achieve higher bacteria removals to act as a cell lysis agent and therefore enhancing both the filter performance and the antifouling properties. 8742



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Figure 3. Bacterial retention mediated by YSZ microtubes in dead-end filtration mode.

(3.2). Microtube Functionalization: Activation Followed by Silanization. Figure 4 presents the hydroxyl group loadings after membrane activation (A) as well as the APTES loadings after silanization (B) where the loading capacities are indicated in OH/nm2 and NH2/nm2, respectively. The number of hydroxyl groups on the microtube surface was determined by potentiometric titration for different hydroxylated microtubes based on the treatment with Piranha solution (RT versus 95 °C), NaOH (95 °C), and in steam atmosphere, respectively (part A of Figure 4). According to Lohbauer37 and Lung,43 acidic hydroxylation of zirconia surfaces using Piranha solution is more effective in comparison to alkaline hydroxylation (NaOH) and hydrothermal treatment (steam atmosphere). In relation to the untreated microtube (0.5 OH/nm2), the hydroxyl surface density of microtubes treated with Piranha solution is increased by a factor of 2.4 to final loading capacities of 1.2 OH/nm2, whereas increased temperatures (95 °C compared to RT) show no significant influence. Compared to the untreated microtube, alkaline hydroxylation using NaOH as well as hydrothermal treatment led only to a marginal (factor 1.4) or rather negligible increase of the hydroxyl group density. To provide a sufficient hydroxyl group content on the membrane surface for subsequent silanization, the microtubes are activated by acidic hydroxy lation using Piranha solution at normal temperature (RT). According to Pasternack44 and Howater,45 APTES functiona lization was carried out in an aqueous solution at increased temperature (75 °C) leading to a cross-linked silane structure with a high amount of available amino groups for subsequent lysozyme immobilization. Here, the incubation time was changed, and tested microtubes were named amino-activated 1 (15 min), amino-activated 2 (90 min), and amino-activated 3 (24 h), respectively. APTES loadings were determined by quantitative acid orange II assay, and part B of Figure 4 shows that it is possible to reach loading capacities between 0.07 and 0.14 NH2 groups per nanometer squared proving the effect of longer incubation times on increased APTES loadings. To ensure a straightforward and highly efficient surface functiona lization procedure providing sufficient terminal amino groups for following lysozyme immobilization, amino-activated type 2 (90 min) with a loading capacity of 0.08 NH2/nm2 is used in the ongoing work.

Figure 4. Loading capacities of hydroxylated (A) and amino-activated (B) YSZ microtubes.

(3.3). Bacterial Adsorption on Functionalized Microtubes. In comparison to the hypothesized lysis of bacteria by lysozyme-functionalized microtubes, the bacterial adsorption behavior of the prefunctionalized microtubes needed for final lysozyme immobilization was investigated. Figure 5 shows the results of bacterial adsorption tests using M. luteus as a function of three different microtube functionalization grades (untreated, hydroxyl-, and amino-activated) as well as three different adsorption times (10 min, 1 h, and 4 h). Bacterial adsorption tests were performed at flow conditions at neutral pH where M. luteus is negatively charged because of its isoelectric point (IEP) of 3.2.46 The tested microtubes with different surface modifications including untreated, hydroxyl-, and amino-activated surface feature IEPs of 6.6, 7.2, and 8.5, respectively (data not shown). Compared to the initial adsorption after 10 min of M. luteus on the untreated and hydroxyl-activated microtube of nearly 8%, the amino-activated microtube showed an increased 8743



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Finally, the lysozyme loading capacity after washing based on the covalent binding (0.0031 ± 0.002 LYS/nm2) is significantly higher (factor 6) compared to the unspecific binding (0.0005 ± 0.001 LYS/nm2). These results make clear that it is possible to convert the inert YSZ microtube as well as the amino-activated microtube into lysozyme-functionalized membranes and these matrices seem to offer high potential for bacteria filtration if a sufficient antibacterial activity can be ascertained. (3.4.2). Lysozyme Activity. To demonstrate the antibacterial behavior of lysozyme-functionalized microtubes, lysozyme activities were analyzed. Thus, agar plate tests for qualitative determination of lysozyme activities (parts A and B of Figure 7) as well as quantitative analysis of lysozyme activities at flow conditions (parts C and D of Figure 7) were performed using gram-positive bacterium M. luteus as substrate. Parts A and B of Figure 7 depict results of the agar plate tests for the lysozyme-functionalized microtubes derived from the unspecific and covalent lysozyme immobilization respectively, and in each case an untreated microtube (no lysozyme immobilization) serves as reference. As expected, bacterial cell growth after incubation for four days is only visible close to the untreated microtubes, whereas both lysozyme-functionalized microtubes based on the unspecific and covalent binding respectively prevent the growth of M. luteus cells. To quantify the lysozyme activities at flow conditions, part C of Figure 7 presents the total values indicated in enzyme units and reduction of CFU/mL/min respectively as a function of the flow rate (0.2−0.5 mL/min). Part D of Figure 7 shows the specific lysozyme activities indicated in enzyme units per milligram of immobilized lysozyme in relation to the applied flow rates. Concerning the total lysozyme activities (part C of Figure 7), two main tendencies turn out as expected. First, an increase of the flow rate leads to a decrease of lysozyme activity because the contact time between the immobilized lysozyme molecules at the membrane surface and the bacteria cells in solution is strongly reduced. On the basis of the used flow rates of 0.2−0.5 mL/min, an increase of the flow rate of 0.1 mL/min leads to a decrease of lysozyme activity of 46−51% and 42−60% for the unspecific and covalent immobilization, respectively. Second, compared to the unspecific binding higher lysozyme activities by a factor of 1.7−2.0 are obtained for the covalent binding, which is in accordance with determined lysozyme loadings based on the results of the Bradford assay (Figure 6). Furthermore, part D of Figure 7 presents the specific lysozyme activities as a function of the flow rate. In contrast to the total lysozyme activities (part C of Figure 7), higher specific enzyme activities by a factor of 3.6−4.1 are obtained for the unspecific lysozyme immobilization (968−96 U/mg) in comparison to the covalent lysozyme immobilization (248− 29 U/mg). Here, the covalent binding of lysozyme to the amino-activated membrane support may lead to conformational changes in enzyme structure, thus, affecting the enzyme’s active site. Combined with the influence of the spacer length on enzyme activities, where in our case APTES was used as relatively short spacer arm with 6 atom length, steric hindrance can result in loss of enzymatic activity.47−49 However, compared to the unspecific binding, lysozyme-functionalized microtubes derived from the covalent functionalization are preferred for generating antibacterial membrane surfaces because of the significant higher total amount of immobilized

Figure 5. Bacterial adsorption at flow conditions (0.2 mL/min) using M. luteus as substrate for different functionalized microtubes.

bacterial adsorption by a factor of 2.5 (around 19%). This can be explained by the different IEPs of tested microtubes where the untreated and hydroxyl-activated samples respectively are only slightly charged at the applied neutral pH conditions, whereas the amino-activated microtube is positively charged acting presumably as a polymer brush for enhanced bacteria adsorption. In relation to the amino-activated microtube, increased adsorption times of 1 and 4 h led to an increased bacterial adsorption by a factor of 1.3 and 1.4 respectively where the amino-functionalized microtube surface seems to be saturated with bacteria cells after 1 h. (3.4). Lysozyme Immobilization: Loading, Activity, and Leaching. (3.4.1). Lysozyme Loading. To analyze the lysozyme loading (lysozyme molecules/nm2) in consideration of washout effects, a Bradford assay was performed, and the results due to the initial loading before washing and the final loading after washing are presented in Figure 6. Here, for an effective removal of unbound and weakly bound lysozyme molecules, the application of five washing steps is needed.

Figure 6. Lysozyme loading capacities in consideration of washout effects based on the unspecific and covalent immobilization strategy (reaction step 3). 8744



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Figure 7. A and B: Agar plate test demonstrating antibacterial activity of lysozyme-functionalized microtubes. C and D: Total and specific enzyme activities of lysozyme-functionalized microtubes at different flow rates.

lysozyme molecules (factor ∼6), which results in increased enzyme activities (factor ∼2). (3.4.3). Lysozyme Leaching. In agreement with applied flow rates for the lysozyme activity tests (parts C and D of Figure 7) Figure 8 presents the amount of leached lysozyme based on a intracapillary rinsing for 1 h with 0.1 M phosphate buffer (pH 7). The initial lysozyme loading capacities for the unspecific and covalent binding were 0.0005 ± 0.001 LYS/nm2 and 0.0031 ± 0.002 LYS/nm2 respectively, that is total amounts of 170.3 ± 25.9 and 678.1 ± 50.1 μg immobilized lysozyme, respectively (Figure 6). For a flow rate of 0.2 mL/min, the amount of leached lysozyme for the covalent (1.2 ± 0.8 μg) compared to the unspecific binding route (4.2 ± 2.0 μg) is significantly lower by a factor of 3.4, which can be explained by the stronger binding of covalent bound lysozyme molecules. As expected, an increase of the flow rate to 0.5 mL/min leads to an increase of washed-out lysozyme amounts by a factor of around 2 because of enhanced shear stress near to the microtube surface. To maintain a constant high level of antibacterial effectiveness for long-term filtration applications (days), regeneration of the microtubes by reimmobilization with lysozyme might therefore be required.

Figure 8. Lysozyme leaching at different flow rates for 1 h for microtubes derived from the unspecific and covalent binding route.

In summary, the combination of the obtained YSZ microtubes with antibacterial enzymes such as lysozyme leads to promising candidates for sustainable bacteria filtration 8745



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(16) Gao, W.; Liang, H.; Ma, J.; Han, M.; Chen, Z-l.; Han, Z-s.; Li, G-b. Membrane fouling control in ultrafiltration technology for drinking water production: A review. Desalination 2011, 272, 1−8. (17) Drews, A. Membrane fouling in membrane bioreactors − Characterisation, contradictions, cause and cures. J. Membr. Sci. 2010, 363, 1−28. (18) Le-Clech, P.; Chen, V.; Fane, T. A. G. Fouling in membrane bioreactors used in wastewater treatment. J. Membr. Sci. 2006, 284, 17−53. (19) Chan, R.; Chen, V. Characterization of protein fouling on membranes: opportunities and challenges. J. Membr. Sci. 2004, 242, 169−188. (20) Dankovish, T. A.; Derek, G. G. Bactericidal paper impregnated with silver nanoparticles for point-of-use water treatment. Environ. Sci. Technol. 2011, 45, 1992−1998. (21) Veerapandian, M.; Yun, K. Functionalization of biomolecules on nanoparticles: Specialized for antibacterial applications. Appl. Microbiol. Biot. 2011, 90, 1655−1667. (22) Parisien, A.; Allain, B.; Zhang, J.; Mandeville, R.; Lan, C. Q. Novel alternatives to antibiotics: Bacteriophages, bacterial cell wall hydrolases, and antimicrobial peptides. J. Appl. Microbiol. 2008, 104, 1−13. (23) Jenssen, H.; Hamill, P.; Hancock, R. E. W. Peptide antimicrobial agents. Clin. Microbiol. Rev. 2006, 19, 491−511. (24) Farnaud, S.; Evans, R. W. Lactoferrin − a multifunctional protein with antimicrobial properties. Mol. Immunol. 2003, 40, 395− 405. (25) Edwards, J. V.; Prevost, N. T.; Condon, B.; French, A. Covalent attachment of lysozyme to cotton/cellulose materials: protein verses solid support activation. Cellulose 2011, 18, 1239−1249. (26) Mueller, B.; Zacharias, M.; Rezwan, K. Bovine serum albumin and lysozyme adsorption on calcium phosphate particles. Adv. Eng. Mater. 2010, 12, B53−B61. (27) Kroll, S.; Treccani, L.; Rezwan, K.; Grathwohl, G. Development and characterisation of functionalised ceramic microtubes for bacteria filtration. J. Membr. Sci. 2010, 365, 447−455. (28) Treccani, L.; Maiwald, M.; Zöllmer, V.; Busse, M.; Grathwohl, G.; Rezwan, K. Antibacterial and abrasion-resistant alumina micropatterns. Adv. Eng. Mater. 2009, 11, B61−B66. (29) Wang, Q.; Fan, X.; Hu, Y.; Yuan, J.; Cui, L.; Wang, P. Antibacterial functionalization of wool fabric via immobilizing lysozymes. Bioproc. Biosyst. Eng. 2009, 32, 633−639. (30) Rezwan, K.; Studart, A. R.; Vörös, J.; Gauckler, L. J. Change of ζ potential of biocompatible colloidal oxide particles upon adsorption of bovine serum albumin and lysozyme. J. Phys. Chem. B 2005, 109, 14469−14474. (31) Senel, S.; Kassab, A.; Arica, Y.; Say, R.; Denizli, A. Comparison of adsorption performances of metal−chelated polyamide hollow fibre membranes in lysozyme separation. Colloids Surf., B 2002, 24, 265− 275. (32) Callewaert, L.; Walmagh, M.; Michiels, C. W.; Lavigne, R. Food applications of bacterial cell wall hydrolases. Curr. Opin. Biotech. 2011, 22, 164−171. (33) Benkerroum, N. Antimicrobial activity of lysozyme with special relevance to milk. Afr. J. Biotechnol. 2008, 7, 4856−4867. (34) Ohlrogge, K., Ebert, K. Membranes − Basics, Proceedings and Industrial Applications, 1st ed.; Wiley-VCH: Weihheim, 2006. (35) Irmler, H. W. Dynamic Filtration with Ceramic Membranes, 1st ed.; Vulkan-Verlag GmbH: Essen, 2001. (36) Garmash, E. P.; Kryuchkov, N. Y.; Pavlikov, V. N. Ceramic membranes for ultra- and microfiltration. Glass Ceram+ 1995, 52, 150−152. (37) Lohbauer, U.; Zipperle, M.; Rischka, K.; Petschelt, A.; Müller, F. A. Hydroxylation of dental zirconia surfaces: Characterization and bonding potential. J. Biomed. Mater. Res. B 2008, 87, 461−467. (38) Fischer, H.; Niedhart, C.; Kaltenborn, N.; Prange, A.; Marx, R.; Niethard, F. U.; Telle, R. Bioactivation of inert alumina ceramics by hydroxylation. Biomaterials 2005, 26, 6151−6157.

compared to commonly used disinfection methods (e.g., chlorine, ozone, UV treatment) because of the possibility of a repeated or continuous reuse of the immobilized enzyme as well as the prevention of toxic or carcinogenic disinfection byproducts. To cover a wide spectrum of ceramic membranes, this straightforward covalent lysozyme immobilization may also be transferred to other ceramic oxide membranes such as alumina or titania membranes for generating antibacterial porous matrices.

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AUTHOR INFORMATION

Corresponding Author

*Phone: +49-421-218-64933, fax: +49-421-218-64932, e-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by German Research Foundation (D.F.G.) within the Research Training Group 1375 “Nonmetallic Porous Structures for Physical-Chemical Functions”.

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REFERENCES

(1) Sutherland, K. Drinking and pure water: Filtration improvements progress global clean water provision. Filtr. Separat. 2012, 49 (12−14), 16. (2) Onda, K.; LoBuglio, J.; Bartram, J. Global access to safe water: Accounting for water quality and the resulting impact on MDG progress. Int. J. Environ. Res. Public Health 2012, 9, 880−894. (3) Frechen, F.-B.; Exler, H.; Romaker, J.; Schier, W. Long-term behaviour of a gravity-driven dead end membrane filtration unit for potable water supply in cases of disasters. Water Sci. Technol. 2011, 11, 39−44. (4) Cabral, J. P. S. Water microbiology. Bacterial pathogens and water. Int. J. Environ. Res. Public Health 2010, 7, 3657−3703. (5) Shannon, M. A.; Paul, W. B.; Elimelech, M.; Georgiadis, J. G.; Marinas, B. J.; Mayes, A. M. Science and technology for water purification in the coming decades. Nature 2008, 452, 301−310. (6) Montgomery, M. A.; Elimelech, M. Water and sanitation in developing countries: Including health in the equation. Environ. Sci. Technol. 2007, 41, 17−24. (7) Nagarnaik, P. M.; Mills, M. A.; Boulanger, B. Concentrations and mass loadings of hormones, alkylphenols, and alkylphenol ethoxylates in healthcare facility wastewaters. Chemosphere 2010, 78, 1056−1062. (8) Tsuno, H.; Kawamura, M. Development of an expanded-bed GAC reactor for anaerobic treatment of terephthalate-containing wastewater. Water Res. 2009, 43, 417−422. (9) Emmanuel, E.; Pierre, M. G.; Perrodin, Y. Groundwater contamination by microbiological and chemical substances released from hospital wastewater: Health risk assessment for drinking water consumers. Environ. Int. 2009, 35, 718−726. (10) Kurniawan, T. A.; Chan, G. Y. S.; Lo, W.-H.; Babel, S. Physicochemical treatment techniques for wastewater laden with heavy metals. Chem. Eng. J. 2006, 118, 83−98. (11) Stevik, T. K.; Aa, K.; Ausland, G.; Hanssen, J. F. Retention and removal of pathogenic bacteria in wastewater percolating through porous media: A review. Water Res. 2004, 38, 1355−1367. (12) Peters, T. Membrane technology for water treatment. Chem. Eng. Technol. 2010, 33, 1233−1240. (13) Van Reis, R.; Zydney, A. Bioprocess membrane technology. J. Membr. Sci. 2007, 297, 16−50. (14) Kentish, S. E.; Stevens, G. W. Innovations in separations technology for the recycling and re-use of liquid waste streams. Chem. Eng. J. 2001, 84, 149−159. (15) Van Reis, R.; Zydney, A. Membrane separations in biotechnology. Curr. Opin. Biotech. 2001, 12, 208−211. 8746



Article

(39) Kroll, S.; Meyer, L.; Graf, A.-M.; Beutel, S.; Glökler, J.; Döring, S.; Klaus, U.; Scheper, T. Heterogeneous surface modification of hollow fiber membranes for use in micro-reactor systems. J. Membr. Sci. 2007, 299, 181−189. (40) Santoso, F. Investigations to the simultaneous amination and pore opening of poly(ether imide) membranes. Ph.D. Dissertation, University of Berlin, Germany, 2004. (41) Bradford, M. M. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 1976, 72, 248−254. (42) Shugar, D. The measurement of lysozyme activity and the ultraviolet inactivation of lysozyme. Biochim. Biophys. Acta 1952, 8, 302− 309. (43) Lung, C. Y. K.; Kukk, E.; Hagerth, T.; Matinlinna, J. P. Surface modification of silica-coated zirconia by chemical treatments. Appl. Surf. Sci. 2010, 257, 1228−1235. (44) Pasternack, R. M.; Amy, S. R.; Chabal, Y. J. Attachment of 3(aminopropyl)-triethoxysilane on silicon oxide surfaces: dependence on solution temperature. Langmuir 2008, 24, 12963−12971. (45) Howarter, J. A.; Youngblood, J. P. Optimization of silica silanization by 3-amino-propyltriethoxysilane. Langmuir 2006, 22, 11142−11147. (46) Martienssen, M.; Reichel, O.; Kohlweyer, U. Surface properties of bacteria from different wastewater treatment plants. Acta Biotechnol. 2001, 21, 207−225. (47) Fang, Y.; Huang, X.-J.; Chen, P.-C.; Xu, Z.-K. Polymer materials for enzyme immobilization and their application in bioreactors. BMB Rep. 2011, 44, 87−95. (48) Sherif, H. E.; Di Martino, S.; Travasio, P.; De Maio, M.; Portaccio, M.; Durante, D.; Rossi, S.; Canciglia, P.; Mita, D. G. Advantages of using non-isothermal bioreactors in agricultural waste water treatment by means of immobilized urease. Study on the influence of spacer length and immobilization method. J. Agric. Food Chem. 2002, 50, 2802−2811. (49) De Maio, A.; El-Masry, M. M.; Portaccio, M.; Diano, N.; Di Martino, S.; Mattei, A.; Bencivenga, U.; Mita, D. G. Influence of the spacer length on the activity of enzymes immobilised on nylon/ polyGMA membranes Part 1. Isothermal conditions. J. Mol. Catal. B: Enzym. 2003, 21, 239−252.

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Highly Efficient Enzyme-Functionalized Porous Zirconia Microtubes for

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