Encapsulation of Bacterial Cells in Electrospun Microtubes

May 12, 2009 - Encapsulation of whole microbial cells in microtubes for use in bioremediation of pollutants in water systems was the main focus of thi...
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Biomacromolecules 2009, 10, 1751–1756

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Encapsulation of Bacterial Cells in Electrospun Microtubes S. Klein,† J. Kuhn,‡ R. Avrahami,§ S. Tarre,† M. Beliavski,† M. Green,† and E. Zussman*,§ Faculty of Civil and Environmental Engineering, Faculty of Biology, and Faculty of Mechanical Engineering, Technion, Israel Institute of Technology Haifa 32000, Israel Received February 8, 2009; Revised Manuscript Received April 21, 2009

Encapsulation of whole microbial cells in microtubes for use in bioremediation of pollutants in water systems was the main focus of this investigation. Coelectrospinning of a core polymeric solution with bacterial cells and a shell polymer solution using a spinneret with two coaxial capillaries resulted in microtubes with porous walls. The ability of the microtube’s structure to support cell attachment and maintain enzymatic activity and proliferation of the encapsulated microbial cells was examined. The results obtained show that the encapsulated cells maintain some of their phosphatase, β-galactosidase and denirification activity and are able to respond to conditions that induce these activities. This study demonstrates electrospun microtubes are a suitable platform for the immobilization of intact microbial cells.

1. Introduction Bioremediation is rapidly becoming an increasingly important tool in reclaiming polluted water resources with which to satisfy the ever growing global water demand. The technologies employed in bioremediation are varied; the most promising of these is based on the immobilization of bacterial cells capable of removing specific contaminants. Cell immobilization techniques have been investigated extensively for numerous applications including biomedical, agricultural systems, and for the bioremediation of pollutants in soils and water. Immobilization of microbial cells has been found to enhance the stability of cell enzymatic activities, protect the cells from mechanical or chemical damage, and sustain large bacterial populations for extended periods.1,2 An example of partial cell immobilization is the controlled growth of biofilms on small diameter carrier particles such as sand or granulated activated carbon (GAC) found in bioreactors for the treatment of wastewater. However, due to the nonsterile conditions typical to these treatment systems, specific microorganisms, such as the atrazine degrading bacteria Pseudomonas sp. strain ADP, grown in such bioreactor systems have suffered from contamination, loss of the ability to degrade atrazine, and eventual washout.3,4 A commonly used whole cell immobilization method that can alleviate the problem of contamination is the entrapment of cells in polysaccharide gels such as alginates, agarose, k-carrageenan, and other polymeric matrices such as gelatin and polyvinyl alcohol.5 Calcium-alginate cross-linking represents an example of a simple and cheap technique that is very suitable for maintaining cell viability due to its relatively mild and nontoxic effects on cells.6 However, this material has a relatively low mechanical strength and the cells entrapped in the Ca-alginate beads leak out and grow in the surrounding medium. Also, the nonhomogenous cell distribution within these beads leads to limitations with regard to diffusion and a reduction in the active surface area.7,8 * To whom correspondence should be addressed. E-mail: meeyal@ tx.technion.ac.il. † Faculty of Civil and Environmental Engineering. ‡ Faculty of Biology. § Faculty of Mechanical Engineering.

In the current work we report a novel method for the immobilization of whole bacterial cell which can be used in bioremediation. Here, the bacteria are encapsulated within electrospun core-shell microtubes. Electrospinning is a commonly used process for generating ultrafine, polymer-based fibers with diameters ranging from tens to hundreds of nanometers.9,10 Electrospun fibers can be produced from a variety of synthetic and natural polymers and are currently being extensively used in biomedical applications such as scaffolds and carriers for biologically active molecules such as proteins and enzymes.11-13 Encapsulation of living cells by electrospinning is of considerable technical interest not only for bioremediation. While encapsulation of cells in nanofibers has been previously achieved, at present, the fabricated fibers are either soluble in water or lose their integrity in aqueous medium. Jaysinghe et al.14 and Townsend et al.15 encapsulated mammalian cells by using a coaxial needle arrangement with the concentrated biosuspension in the inner needle and medical grade polydimethylsiloxane (PDMS) in the outer needle. The post electrospinning cells were found to be viable without cellular damage. Salalha et al.16 described the encapsulation of whole bacterial cells in electrospun polyvinyl alcohol (PVA) nanofibers. There it was shown that bacterial cells and viruses can survive the electrospinning process and that these retain their viability even after storing them dry as mats for a number of months. Gensheimer et al.,17 on the other hand, found considerable variability in bacterial viability after producing electrospun polyethylene oxide (PEO) nanofibers. Although demonstrating their potential for live cell encapsulation, the existing electrospun fibers are of limited use for bioremediation. To increase the applicability of live cell encapsulation by electrospinning, fibers are needed that are insoluble in water and are not biodegradable or biodegradable for a predetermined time period. Furthermore, immobilization of microbial cells in a confined space such as a solid fiber may negatively affect their behavior as suggested by the findings from the immobilization of microbial cells in sol-gels.18 The development of electrospun hollow polymeric microfibers (microtubes) offers a new method for bacterial cell encapsulation.19 In this single-step coelectrospinning process, the potentially toxic organic phase consists of a water-insoluble polymer (i.e., the outer shell solution) that is separated from the aqueous

10.1021/bm900168v CCC: $40.75  2009 American Chemical Society Published on Web 05/12/2009

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phase (i.e., the core solution). It was shown that proper selection of the polymers and solvents results in hollow polymeric microfibers. The fabricated microtubes are potentially beneficial to microbial cells by protecting them while providing space for the cells to divide and accumulate. This technology has been successfully applied for the encapsulation of pure enzymes.20 In that study, the addition of polyethylene glycol (PEG) to the shell solution altered the shell’s morphology and made it more porous which thereby positively affected the transfer of small molecules into and out of the microtubes. In the present investigation we demonstrate the feasibility of as-spun microtubes as a method for the immobilization of whole bacterial cells.21 This encapsulation approach has a potentially wide range of applications particularly for use in aqueous environments which includes the biodegradation of hazardous materials, bioremediation of pollutants in water systems, and bioconversions in the biotechnology industry.

2. Materials and Methods 2.1. Bacterial Strains and Growth Conditions. Three different species of bacteria were used in these studies. An Escherichia coli strain that can grow on lactose and is also resistant to the antibiotic kanamycin was examined when β-galactosidase was measured. Pseudomonas ADP is able to grow with the herbicide atrazine as the sole source of nitrogen and synthesizes an alkaline phosphatase whose activity can be measured in whole cells. The third strain was from Pseudomonas putida into which a gene specifying the red fluorescent protein had been introduced (see Supporting Information for details of its construction) and this strain was used in conjunction with confocal microscopy when cells in microfibers were examined. In Escherichia coli the synthesis of β-galactosidase, which normally splits lactose to galactose and glucose, can be repressed by the addition of glucose to the medium while, when glucose is lacking, the enzyme is synthesized in large amounts in the presence of isopropyl-thiogalactoside (IPTG), a synthetic inducer that is not cleaved by this enzyme. Pseudomonas ADP was propagated at 30 °C in a MOPS based medium containing citrate as the carbon source (see Supporting Information) with excess orthophosphate to suppress the synthesis of alkaline phosphatase or with limiting phosphate to induce this enzyme. For denitrification assays, the strain was initially grown aerobically with atrazine present and then transferred to anaerobic conditions in which nitrite was the sole source of nitrogen. Pseudomonas putida S12::dsred was grown at 30 °C in the minimal salts medium mentioned above with the addition of kanamycin (30 mg/L) to maintain the added genetic element containing the gene encoding the red fluorescent protein. Doubly distilled water (DDW) was used throughout. Microscopic images of the distribution of P. putida S12::dsred cells inside the microtubes were obtained with cells grown to stationary phase with shaking at 30 °C in 2 mL of the media cited above. Cells were harvested by centrifugation, the pellets washed twice in DDW and then resuspended with 5% glycerol to a total volume of 50 µL. This was mixed with 450 µL of core solution. The bacterial concentration used for electrospinning corresponded to an optical density at 600 nm [A600] of about 2 as measured with a UVmini-1240 spectrophotometer (Shimadzu, Kyoto, Japan). E. coli KT4/RP4 cells were grown to stationary phase by shaking overnight at 37 °C in 10 mL of LB-kanamycin-IPTG medium. Cells were harvested by centrifugation, the pellets washed with 5 mL of Z buffer (0.1 M sodium phosphate (pH 7.0), 10 mM KCl, 1 mM MgSO4, 5 mM mercaptoethanol) and then resuspended with 1 mL of Z buffer. For electrospinning, 40 µL of these concentrated cells were mixed with 360 µL of core solution. The bacterial concentration used for electrospinning was 3.8 × 109 CFU/mL. Pseudomonas ADP cells were grown to stationary phase with shaking at 30 °C in 8 mL of MOPS medium and were either starved for

Klein et al. phosphate (0.1 mM of K2HPO4) or with excess phosphate (2 mM of K2HPO4). Cells were harvested by centrifugation and the resulting pellets were washed twice with water. Cells were resuspended with 5% glycerol to a total volume of 50 µL and then mixed with 450 µL of core solution. The bacterial concentration used for electrospinning and free cell suspension assays corresponded to an optical density at 600 nm [A600] of about 10. Pseudomonas ADP cells were grown to stationary phase with shaking at 30 °C in 10 mL of a salt-based medium containing nitrite. Cells were harvested by centrifugation and the resulting pellets were washed twice with water. Cells were resuspended with 5% glycerol to a total volume of 100 µL and then mixed with 900 µL of core solution. The bacterial concentration used for electrospinning and free cell suspension assays corresponded to an optical density at 600 nm [A600] of about 12. 2.2. Electrospinning. Preparation of Polymer Solutions. The shell solution was 9 wt % polycaprolactone (PCL) 80 K + 1 wt % polyethylene glycol (PEG) 6 K dissolved in a mixture of chloroform and dimethylformamide (DMF), 90:10 (w/w); the core solution was 5 wt % polyethylene oxide (PEO) 600 K in H2O. All polymers were purchased from Sigma-Aldrich and were used as is. Electrospinning Set Up. Core-shell fibers were fabricated by a coelectrospinning process using the set up described by Dror et al.19 All experiments were conducted at room temperature (∼23 °C) and a relative humidity of about 40%. The spinning parameters were as follows: the electrostatic field used was approximately 0.7 kV/cm and the distance between the spinneret and collector plate was 14 cm. The flow rates of both the core and shell solutions were controlled by two syringe pumps and were 3.5 mL/h for the shell and 0.5 mL/h for the core. The fibers were collected as a strip on the edge of a vertical rotating wheel22 having a velocity of 1.2 m/sec. For fluorescence microscopy, a few fibers were collected directly onto a microscope slide. 2.3. Measurement of Enzyme Activity. β-Galactosidase ActiVity. For the assay encapsulated cells, whole mats were weighed directly after electrospinning and cut up into pieces that weigh 20-30 mg each. Following several washes in Z buffer, the pieces where dipped in 1.0 mL of Z buffer for 60 min. The reaction23 was started by the addition of 0.2 mL of orthonitrophenyl-β-D-galactoside (ONPG) at a concentration of 4 mg/mL. Incubation was at 37 °C until the appearance of the reaction’s yellow product, nitrophenol, became visible and then the reaction was stopped by adding 0.5 mL of 1 M Na2CO3. A total of 1 mL of the assay mixture was transferred to a spectrophotometer cuvette. For the assay of free cells, 20 µL of the same cell suspension prepared for electrospinning was added to 0.98 mL of Z buffer. The cells were allowed to equilibrate for 30 min at 37 °C before the addition of the substrate. The reaction was conducted in the same manner as that for encapsulated cells except for an additional step after stopping the reaction which removes cells from the assay mixture by centrifugation. Assay mixtures, including blanks (reaction without cells), were read at 410 nm (Genesys 10 UV, Thermo Scientific, MA, U.S.A.). Enzyme activity was computed in Miller Units (MU; the change in absorbance of light per cm at 410 nm (A410) multiplied by 1000 divided by the measured reaction time in minutes (t)) and either multiplied by the dilution used (free cell suspension) or divided by the fraction of the electrospun mat weight sampled. Phosphatase ActiVity. For the assay of encapsulated cells, whole mats were weighed directly after electrospinning and then cut up into pieces that weighed 20-30 mg each. Following several washes in 50 mM Tris HCl-Trizma (pH 7.3), the pieces where immersed in 1.4 mL of the same Tris buffer for 60 min. To start the reaction, 0.6 mL of p-nitrophenyl phosphate (7 mg/mL in H2O; Calbiochem Corp.) was added. The reaction was performed at 25 °C until the appearance of the yellow product, nitrophenol. The reaction was stopped by adding 0.5 mL of 1 M Na2CO3. A total of 1 mL of the assay mixture was transferred to a spectrophotometer cuvette. For the assay of free cells

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Figure 1. (a) HRSEM micrographs of microtubes used for encapsulating living cells; (b,c) fluorescent microscopy images of encapsulated P. putida S12::dsred.

in suspension, the cells were prepared in the same manner as that for cells prepared for electrospinning with the exception that the pellets after washing were resupended with 5% glycerol to a total volume of 50 µL and then mixed with 0.45 mL 0.85% NaCl. A 100 µL aliquot of this free cell suspension was added to 0.6 mL of 50 mM Tris HClTrizma (pH 7.3). The cells were allowed to equilibrate for 30 min at 25 °C before addition of 0.3 mL substrate. The reaction was conducted in the same manner as for encapsulated cells except for the additional step of removing the cells from the assay mixture by centrifugation. Assay mixtures and blanks (reaction without cells) were read at 410 nm (Genesys 10 UV, Thermo Scientific, MA, U.S.A.). Enzyme activity was computed as phosphatase units (arbitrary), which were calculated in the same manner as that for Miller Units with β-galactosidase activity. Denitrification ActiVity. For both respiration and induction experiments, denitrification activity was measured as the disappearance of nitrite from the growth medium. For assays of encapsulated cells, whole mats were weighed directly after electrospinning and cut up into pieces that weigh about 200 mg each. Each piece was further divided into eight small pieces. Following several washes with water, the pieces were placed in a 15 mL tube filled to the top with only minimal salt based medium and left at room temperature for 18 h. At the end of that period, the medium was discarded and the pieces were washed twice using a vortex to wash away any cells that might be on the exterior of the microtubes. The pieces were then transferred to a small vial filled with 4 mL of minimal salt base medium supplemented with trisodium citrate at a concentration of 2 g/L and NaNO2- at concentration of 25 mg/L as the nitrogen source. The medium was then bubbled for 40 min with nitrogen gas to remove dissolved oxygen. At the end of the nitrogen bubbling period, the first sample (t ) 0) was taken and the vial was sealed. Samples were subsequently taken at different times while nitrogen gas was bubbled through the medium to maintain anoxic conditions. Samples (100 µL) were diluted immediately with water and analyzed for nitrite. Nitrite concentration was determined by absorption at 543 nm (UVmini-1240, Shimadzu, Kyoto, Japan) using the sulfanilamide colorimetric method.24 For suspensions of free cells, assays of both cells grown under denitrifying or aerobic conditions were prepared in the same manner as that outlined for electrospinning with the exception that the pellets, after washing, were resuspended with 5% glycerol to a total volume of 100 µL and then mixed with 0.9 mL

0.85% NaCl. The assay was conducted in the same manner as that for encapsulated cells with two exceptions: (1) To keep the same ratio of bacteria to medium used in the encapsulated cells assay, the assay volume was 12 mL; (2) Samples (100 µL to 1 mL) were diluted with DDW and filtered with 0.45 µm membrane filters (Millipore) to remove the cells before being analyzed for nitrite. 2.4. Imaging. Images of the fibers were obtained using a Leo Gemini high resolution scanning electron microscope (HRSEM) at an acceleration voltage of 3 kV and a sample to detector distance of 3-5 mm. The specimens were coated with a thin gold film to increase their conductivity. The distribution of encapsulated P. putida S12::dsred cells inside the microtubes was visualized by a Zeiss Axiovert 200 M microscope (Go¨ttingen, Germany), using filters for detection of DsRed (excitation/emission: 545 nm ( 25/605 nm ( 70). Images were captured using Hdcam (Hamamtsu) 1394 ORCA-ERA and processed with AxioVision LE 4.5 software.

3. Results and Discussion 3.1. Microtubes Morphology and Cell Distribution within the Microtubes. Core-shell fibers were fabricated with the polymeric solutions described in the Experimental Section. The resultant fibers are partially collapsed microtubes with porous walls which are due to PEG in the shell solution. Cross sections of the microtubes and their surface morphology are shown in Figure 1a. As previously shown by Arinstein et al.,25,26 the radial collapse of the microtubes is expected for this combination of core-shell solutions. This collapse can occur to such an extent that the microtubes are transformed into a ribbon like structures. The tubular space available for the encapsulated bacteria seems rather restricted, averaging a diameter of 3.5 µm. To observe the location of cells within the electrospun fibers, a Pseudomonas putida strain that was modified to express the red fluorescent protein (DsRed) was encapsulated within the microtubes. Images of the encapsulated fluorescent cells were taken after immersing the slides carrying nonwoven microtubes in water for 2 h to wash away any cells that might be on the

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Table 1. Enzymatic Activity of Encapsulated Whole Cells enzyme

species

source

enzyme activity unitsa

relative activityb (%)

phosphatase(s)

Pseudomonas ADP

β-galactosidase

E. coli

electrospun mats 8 h in bufferd 24 h bufferd electrospun mats

124.1 ( 32.3 2.9 ( 0.6 3.1 ( 0.3 15153.7 ( 566.5

9.8 0.2/2.3c 0.2/2.5c 22.8

a

Phosphatase activity: arbitrary units (PU) that are given as the change in absorbance at a wavelength of 410 nm (A410) multiplied by 1000 and divided by the time in minutes. This number is then multiplied by either the dilution used (for free cell suspension) or the weight of the entire mat divided by the weight of the sampled piece. β-galactosidase activity: measured as Miller Units (MU) that are calculated in the same manner as phosphatase units. b Free cells versus encapsulated cells. Phosphatase activity (PU) of free cells: 1270.88 ( 109.0. β-galactosidase activity of free cells: 66376.67 ( 7998.5 MU. c Relative activity of electrospun microbial cells. d The buffer samples measure the leakage of phosphatase from the microtubes.

exterior of the microtubes. Figure 1b,c shows the location of encapsulated fluorescent cells within the electrospun microtubes. The encapsulated cells appear to be aligned with and unevenly distributed along the microtube axis. At the concentration of cells used (109), the microtubes appear to be sparsely populated. 3.2. Enzymatic Activity of Encapsulated Whole Cells. The impact of electrospinning on bacterial cells was initially tested on their enzymatic activity after encapsulation. Two enzymes were selected, alkaline phosphatase and β-galactosidase. The assays were conducted on two different bacterial species that had been encapsulated in the same manner. The results are shown in Table 1. The enzymatic activity of the encapsulated cells within a sample of fibers was compared to that of free cell suspensions (before electrospinning). To induce phosphatase synthesis, Pseudomonas ADP cells were grown under phosphate limiting conditions. Phosphatase activity was indeed found in the encapsulated Pseudomonas ADP cells grown in this way. However, the microtubes contained only about 10% activity of that of the free cells prior to electrospinning. To determine whether this enzyme diffused out of the fibers as a result of cell disruption during electrospinning, phosphatase activity was also measured outside the nonwoven mats. The assay was conducted on samples excised from the same nonwoven mats used for the phosphatase activity assay. After an initial rinsing of them, the excised pieces were placed in buffer for 24 h. Samples from the buffer were assayed for enzymatic activity after 8 and 24 h. As shown in the Table 1, the relative activity found in the buffer was very low and did not change over time. This is in contrast to recent findings with encapsulated purified alkaline phosphatase that showed that there is significant leakage from these fibers to the buffer in the first 24 h.20 Results from the current phosphatase experiment suggest that the encapsulated cells maintained their membrane integrity and leakage of intercellular enzymes is thereby minimal. Dror et al.20 suggested that the relatively small size (80 kDa) of alkaline phosphatase accounted for the massive leakage observed in their experiments because a similar experiment with a larger size enzyme, β-galactosidase (465 kDa), showed only a small amount of enzyme detected in the buffer. The second enzymatic activity examined was β-galactosidase. Encapsulated E. coli KT4/RP4 cells that had been grown with isopropyl-thiogalactoside (IPTG) prior to electrospinning were assayed. In this case, 23% of the enzymatic activity remained after electrospinning. The slightly better recovery of the enzymatic activity might be related to a greater durability of this bacterial species or of this enzyme during the electrospinning process. The enzymatic activity observed for both enzymes does not depend on the survival of viable cells after the electrospinning process. Even though the electrospinning process negatively affected enzymatic activity as compared to free cells, the assays results were an encouraging first step because they point to the potential of electrospinning as a platform for encapsulation

enzymes in whole cells. The low recovery of enzymatic activity suggests the addition of a “microtubes filling” step following electrospinning would be very useful. That is, the cells that remain viable can be subsequently grown inside the tubes by immersing them in suitable media. This has been done and those experiments will be the subject of a future communication. 3.3. Encapsulated Cell Viability. Determining cell viability directly is difficult for encapsulated microbial cells in electrospun core-shell fibers. Enzymatic activity reflects cellularly bound enzyme(s) that remain active regardless of whether the cells are alive or dead. Due to the insoluble character of the microtube, a direct assessment of cell viability by the usual platecounting technique is not possible. The physiological properties of encapsulated cells were, therefore, evaluated by cell respiration and their ability to synthesize proteins. Both these properties rely on the membrane integrity of the cells and employ complex systems. Respiration of Encapsulated Whole Cells. Respiration under denitrifying conditions was measured as nitrite disappearance from the growth medium by encapsulated cells and their equivalent free cells suspensions (Figure 2a). As expected, nitrite utilization by a suspension of free cells began immediately and all external nitrite was consumed within 5 h. Encapsulated cells exhibited about a 9 h delay before measurable nitrite uptake began. Complete nitrite utilization was observed after an additional period of about 11 h. The different kinetics of the encapsulated cells as compared to the free cell suspension may be due to (1) mass transfer limitation of nitrite into the microtubes; (2) impact of the electrospinning process on the encapsulated cells; and (3) adaptation of encapsulated cells to their new environment. Because previous results showed a large reduction of enzymatic activity due to the electrospinning process, it is more likely that the number of physiologically active encapsulated cells was insufficient for detecting nitrite utilization at the beginning of the experiment. A subsequent recovery time or growth period was required before nitrite respiration was significant enough to be measured. De NoVo Synthesis of Enzymes in Encapsulated Cells. In the first set of experiments, encapsulated cells were exposed to medium that induces the synthesis of phosphatase. Pieces of the mat containing Pseudomonas ADP cells, previously incubated in MOPS medium containing an excess of phosphate, were placed in growth medium containing a limiting amount of orthophosphate (phosphatase inducing conditions) for 4 days. As expected, phosphatase activity in electrospun mats containing cells that had been grown with a relatively high concentration of orthophosphate was low and averaged 0.146 ( 0.055 PU/20 mg of electrospun mat versus cells grown in low phosphate which gave 6.71 ( 2.4 PU/20 mg. After incubating pieces of this mat in a growth medium containing a limiting amount of orthophosphate, a high level of phosphatase was observed which averaged 248 ( 34 APU/20 mg of electrospun mat. The average increase in phosphatase activity was about 1700 times. This

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A second set of experiments examined the induction of denitrifying activity in encapsulated whole cells. Induction of denitrification activity was based on the strain’s ability to grow under either aerobic or denitrifying (anaerobic) conditions where oxygen or nitrite serve as the terminal electron acceptor, respectively. In general, the main exogenous signals that induce the synthesis of the denitrification system are a low oxygen tension and the presence of an N-oxide that can be used in respiration.27 As shown in Figure 2b, when aerobically grown free cells were transferred to denitrifying growth conditions, they exhibited about an 8 h adaption period (lag time) until denitrifying activity became observable. As previously stated above, cells already grown under denitrifying conditions exhibited relatively high denitrifying activity without any lag time, while the same encapsulated cells had a lag time of 9 h. When encapsulated Pseudomonas ADP cells previously grown under aerobic growth conditions were transferred to denitrifying growth conditions, a much longer lag time of about 15 h was required before denitrifying activity was initiated. At the very least, these results clearly demonstrate that the encapsulated cells respond to an environment that induces synthesis of nitrite reductase.

4. Conclusions

Figure 2. Nitrite utilization by encapsulated whole cells and free cells. (a) Percentage of nitrite remaining in the growth medium with denitrifying free cells (0) and encapsulated denitrifying whole cells (b), and (b) percentage of nitrite remaining in the growth medium with denitrifying free cells (O); aerobically grown free cells (9) and encapsulated aerobically grown whole cells (4).

clearly shows that the encapsulated cells within the electrospun mat were able to respond to the surrounding environment and synthesize phosphatase. In addition, the level of phosphatase activity reached was statistically significantly higher (p < 0.05) than free cells grown under the same limiting phosphate conditions 16.90 ( 1.3 PU and suggests that the encapsulated cells in the microtubes had indeed divided. That is, if the encapsulated cells had only synthesized phosphatase without growing and dividing, then the highest level of phosphatase reached should have been that of the free cells previously grown in low levels of orthophosphate. It is important to note that the large standard deviation observed reflects variation in the impact of electrospinning on cells. It appears that large numbers of cells are killed or incapacitated by the process, with the number of physiologically active cells being 1-2 orders of magnitude less than the amount prior to electrospinning. Gensheirmer et al.17 suggested that the rapid evaporation upon electrospinning may cause a drastic change in the osmotic environment leading to a decrease in cell viability. However, evaporation in the water-based core of microtubes was found by Arinstein et al.25 to be very slow, as evidenced by the buckling effect of the microtubes. The low survival rates observed in the microtubes are probably due to the shear and tension stresses that act on these cells.16

The present study demonstrates the feasibility of novel electrospun microtubes as a method to immobilize bacterial cells. The ability of the microtube structure to support cell attachment and maintain enzymatic activity of the encapsulated microbial cells was examined. This preliminary investigation shows that the encapsulated cells maintain some of their phosphatase, β-galactosidase and denitrification activity and are able to respond to inducing conditions in their surrounding environment. The low number of viable cells that remain recovered their enzymatic activity when the relevant growth conditions were provided. The positive results obtained with encapsulated Pseudomonas ADP, a species capable of degrading the herbicide atrazine are encouraging. This is a first step toward the development cell-bearing microtubes for bioremediation. Bacterial cell encapsulation combined with the ability to control mass transfer through the microtube shell creates a bioreactor like structure. In addition, this fabrication method enables encapsulation of different kinds of bacterial cells in the same microtube. This technique has the unique benefit of a large ratio of surface to volume and should find use when separation between bacterial cells and the external aqueous environment is desired (such as in water purification processes). The as-spun microtubes can be easily integrated into water purification systems and should be able to cover standard biofilm carrier surfaces, such as sand, plastic, and rocks. Microtube technology also offers significant advantages for studying the behavior of cells under confinement. Cells, either bacterial or mammalian, were found to alter their behavior in a confined environment.28-30 The flexibility of the microtube system that permits the control of permeability, fiber diameter (i.e., degree of confinement), and the ability to confine and track individual cells should provide a platform for this type of research. Acknowledgment. We wish to thank the RBNI (Russell Berrie Nanotechnology Institute) and the GWRI (Grand Water Research Institute) at the Technion for supporting this research. Supporting Information Available. Bacterial strains, growth conditions, and DNA manipulation. This material is available free of charge via the Internet at http://pubs.acs.org.

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References and Notes (1) Lee, C. M.; Lu, C. J.; Chuang, M. S. Water Sci. Technol. 1994, 30, 87–90. (2) Adam, W.; Lukacs, Z.; Kahle, C.; Saha-Moller, C. R.; Schreier, P. J. Mol. Catal. B: Enzym. 2001, 11, 377–385. (3) Katz, I.; Dosoretz, C. G.; Mandelbaum, R. T.; Green, M. Water Res. 2001, 35, 3272–3275. (4) Herzberg, M.; Dosoretz, C. G.; Tarre, S.; Beliavski, M.; Green, M. Water Sci. Technol. 2004, 49, 215–222. (5) Cohen, Y. Bioresour. Technol. 2001, 77, 257–274. (6) Anisha, G. S.; Prema, P. Bioresour. Technol. 2008, 99, 3325–3330. (7) Kourkoutas, Y.; Bekatorou, A.; Banat, I. M.; Marchant, R.; Koutinas, A. A. Food Microbiol. 2004, 21, 377–397. (8) Zohar-Perez, C.; Chet, I.; Nussinovitch, A. Biotechnol. Bioeng. 2004, 88, 671–674. (9) Reneker, D. H.; Yarin, A. L.; Zussman, E.; Xu, H. AdV. Appl. Mech. 2007, 41, 43–195. (10) Reneker, D. H.; Fong, H. Polymeric Nanofibers; Oxford University Press: New York, 2005. (11) Ramakrishna, S.; Fujihara, K.; Teo, W. E.; Yong, T.; Ma, Z. W.; Ramaseshan, R. Mater. Today 2006, 9, 40–50. (12) Agarwal, S.; Wendorff, J. H.; Greiner, A. Polymer 2008, 49, 5603– 5621. (13) Greiner, A.; Wendorff, J. H.; Yarin, A. L.; Zussman, E. Appl. Microbiol. Biotechnol. 2006, 71, 387–393. (14) Jayasinghe, S. N.; Irvine, S.; McEwan, J. R. Nanomedicine 2007, 2, 555–567. (15) Townsend-Nicholson, A.; Jayasinghe, S. N. Biomacromolecules 2006, 7, 3364–3369.

Klein et al. (16) Salalha, W.; Kuhn, J.; Dror, Y.; Zussman, E. Nanotechnology 2006, 17, 4675–4681. (17) Gensheimer, M.; Becker, M.; Brandis-Heep, A.; Wendorff, J. H.; Thauer, R. K.; Greiner, A. AdV. Mater. 2007, 19, 2480–2482. (18) Nassif, N.; Bouvet, O.; Rager, M. N.; Roux, C.; Coradin, T.; Livage, J. Nat. Mater. 2002, 1, 42–44. (19) Dror, Y.; Salalha, W.; Avrahami, R.; Zussman, E.; Yarin, A. L.; Dersch, R.; Greiner, A.; Wendorff, J. H. Small 2007, 3, 1064–1073. (20) Dror, Y.; Kuhn, J.; Avrahami, R.; Zussman, E. Macromolecules 2008, 41, 4187–4192. (21) Pieper, D. H.; Reineke, W. Curr. Opin. Biotechnol. 2000, 11, 262– 270. (22) Theron, A.; Zussman, E.; Yarin, A. L. Nanotechnology 2001, 12, 384– 390. (23) Miller, J. H. Experiments in Molecular Genetics; Cold Spring Harbor Laboratory: Cold Spring Harbor, NY, 1972. (24) Standard Methods for the Examination of Water and Wastewater, 19th ed.; American Public Health Association/American Water Works Association/Water Environment Federation: Washington, DC, 1995. (25) Arinstein, A.; Avrahami, R.; Zussman, E. J. Phys. D: Appl. Phys. 2009, 42 (1), 015507. (26) Arinstein, A.; Zussman, E. Phys. ReV. E 2007, 76 (5), 056303. (27) Zumft, W. G. Microbiol. Mol. Biol. ReV. 1997, 61, 533–616. (28) Stamenovic, D.; Ingber, D. E. Soft Matter 2009, 5, 1137–1145. (29) Wang, J. H. C.; Thampatty, B. P. Biomech. Model. Mechanobiol. 2006, 5, 1–16. (30) Nassif, N.; Roux, C.; Coradin, T.; Bouvet, O. M. M.; Livage, J. J. Mater. Chem. 2004, 14, 2264–2268.

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