Living Composites of Electrospun Yeast Cells for Bioremediation and

Sep 9, 2015 - The Faculty of Mechanical Engineering, Technion Institute of Technology, Haifa 32000 Israel. § Macromolecular Chemistry II and Bayreuth...
2 downloads 0 Views 8MB Size
Download PDF
Article pubs.acs.org/Biomac

Living Composites of Electrospun Yeast Cells for Bioremediation and Ethanol Production Ilya Letnik,*,† Ron Avrahami,‡ J. Stefan Rokem,*,† Andreas Greiner,§ Eyal Zussman,‡ and Charles Greenblatt*,† †

Department of Microbiology and Molecular Genetics, IMRIC Hebrew University − Hadassah Medical School Ein-Karem, Jerusalem 9112102, Israel ‡ The Faculty of Mechanical Engineering, Technion Institute of Technology, Haifa 32000 Israel § Macromolecular Chemistry II and Bayreuth Center for Colloids and Interfaces, University of Bayreuth, Universitätsstrasse 30, 95440 Bayreuth, Germany ABSTRACT: The preparation of composites of living functional cells and polymers is a major challenge. We have fabricated such “living composites” by preparation of polymeric microtubes that entrap yeast cells. Our approach was the process of coaxial electrospinning in which a core containing the yeast was “spun” within a shell of nonbiodegradable polymer. We utilized the yeast Candida tropicalis, which was isolated from olive water waste. It is particularly useful since it degrades phenol and other natural polyphenols, and it is capable of accumulating ethanol. The electrospun yeast cells showed significant activity of bioremediation of phenol and produced ethanol, and, in addition, the metabolic processes remained active for a prolonged period. Comparison of electrospun cells to planktonic cells showed decreased cell activity; however, the olive water waste after treatment by the yeast was no longer toxic for Escherichia coli, suggesting that detoxification and prolonged viability and activity may outweigh the reduction of efficiency.

■

INTRODUCTION Immobilization of cells of various kinds has several advantages as compared to the use of free cells.1 Productivity can be increased with immobilization since the cells are usually stable over a longer period of time without losing activity, and this also allows for their reuse.1−3 One of the methods of immobilization is encapsulation in polymer matrices. In this process, the biocatalyst is restricted by a semipermeable polymer, which allows for free flow of substrates and nutrients in, and waste out. Results from bioreactor studies with encapsulated cells have demonstrated advantages over free cells under various conditions. For example, increased and prolonged metabolic activity and metabolite production,4 protection from toxic substances, and also increased plasmid stability have been observed.1,5,6 Polymer fibers produced by electrospinning where shown to be promising agents for whole cell immobilization.7−9 Traditionally, the fibers were made from one polymer; however, a novel modification of the process entails the creation of a twolayered fiber via the process of coelectrospinning.10 Electrospinning is a versatile method that allows the creation of continuous polymer fibers with micro- or nanoscale diameters. In a typical electrospinning experiment, a polymer solution is pumped through a thin nozzle, which simultaneously serves as an electrode, to which a high electric field of 100−500 kVm is applied. In coaxial electrospinning, two concentrically aligned nozzles are used for spinning. The same voltage is applied to both nozzles, and it deforms the compound droplet. A jet is © XXXX American Chemical Society

generated on the tip of the deformed droplet, and a core−shell nanofiber is created. It is possible to produce structures ranging from single fibers to ordered arrangements. The scope of applications is very broad touching on fields as diverse as optoelectronics, sensor technology, catalysis, filtration, and medicine.11 This method suggested itself since (1) fibers consisting of water-soluble polymers such as poly(vinyl alcohol) (PVA) cannot be used in aqueous environments, as in wastewater treatment or any water associated biotechnological application, (2) the coaxial method allows for the creation of a more natural environment for the immobilized cells, due to the water-soluble core polymer, and (3) adjustment of the quality of the shell polymers allows for greater physical durability and flexibility. As far as is known, microbial electrospun cells can survive and maintain their protein conformation for months.8 Yeast cells were electrospun recently and were shown to survive the process and maintain membrane integrity and viability over time.7 In our laboratory, a yeast identified as Candida tropicalis was isolated from industrial olive mill wastewater (OMW). The ability of C. tropicalis to degrade a wide range of environmental pollutants such as phenol, polyphenols, 4-chlorophenol, and even crude oil has been described by different researchers.12−14 Received: July 19, 2015 Revised: September 6, 2015

A

DOI: 10.1021/acs.biomac.5b00970 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

Figure 1. Structure and composition of the polymer tubes. (A) XHR SEM image of a tube’s surface. (B) Shell polymer surface topography and pore sizes. (C) SEM image of a single C. tropicalis cell. (D) Bright field image of several yeasts containing microtubes.

■

In addition, C. tropicalis has high resistance to metals such as chromium and copper and is able to produce xylitol,15,16 all of which can be future possible uses of this species. Here we report the encapsulation of living C. tropicalis cells in microtubes by coaxial electrospinning. The immobilized yeast was characterized for survival and phenol degradation. Phenols are present in many wastewaters from various industries such as petroleum, pulp and paper, pharmaceutical, and wood-processing chemicals. We focused on OMW, which is a liquid byproduct of olive oil production. Olive oil annual production in the Near East and Mediterranean countries reached 2600 kilotons in 2009. 17 One ton of olives approximately produces 0.8 tons of OMW. The high polluting potential of OMW is linked to its low pH (4−5), and the significant amounts of organic molecules, especially polyphenolic mixtures, present with varying molecular weights (2− 10 g/L phenols).18 Wastewaters contaminated by phenols endangers fish life at relatively low concentration, e.g., 5 to 25 mg·L−1.19 The phytotoxic properties of OMW were also shown with spinach as the test object.20 For the treatment of OMW, valorization, reuse, and other various management methods have been proposed (physical, physicochemical, biological and thermal). However, no treatment method is efficient when solely used, since high application costs are involved.17 Thus, reducing the toxicity of OMW by simple biodegradation of the phenolic fraction could provide an effective tool that will aid in environmental protection. Another function of immobilized cells tested was ethanol fermentation, done as a model for metabolite production. In addition, we show that treatment of OMW by the immobilized yeast, reduces its toxicity to Escherichia coli. The extension of this model to “good soil bacteria” will be of interest. In the future, electrospun yeast may be used in bioremediation and other aqueous environment applications as well as in metabolite formation.

MATERIALS AND METHODS

Yeast Strains, Culture Conditions. The C. tropicalis used in this study was isolated from OMW and identified by 16S rDNA sequenced by Hy Laboratories Ltd., Israel. Saccharomyces cerevisiae was obtained from a local commercial supplier. Experiments were done in batch cultures in which the yeast was grown overnight with shaking at 150 rpm at 30 °C. The growth medium was either CASO bullion agar (Carl Roth Gmbh # x938 ref) or minimal medium (3 g/L NaNO3, 1 g K2HPO4, 0.5 g MgSO4, 0.5 g KCl, 0.1 g FeSO4 and 1.5% Agar + phenol (30−100 mg/L) as a single carbon source) . For the encapsulation, the cells were grown in liquid CASO medium for 24 h (to stationary phase) and collected by centrifugation. Cell Immobilization. A core−shell electrospinning technique was used for immobilization of the yeast. The system allows for fabrication of microtubes in an electrostatic field. Two separate syringes continuously ejected solutions from a spinneret with two coaxial capillaries under high voltage. The internal capillary of this spinneret facilitated the flow of the yeast cells in an aqueous solution, while being enveloped by the external capillary containing a nonbiodegradable polymer solution. The core consisted of 20 vol % cell mass in 15 wt % polyvinylpyrrolidone (PVP) Mw = 1300k in deionized water. The shell copolymer consisted of 15% poly vinylidene fluoride-cohexafluoropropylene (PVDF-HFP) Mw = 400k and 2 wt % PEG (M w = 6k) in a solvent mixture of tetrahydrofuran and dimethylformamide (THF/DMF) in a weight ratio of 6.5:3.5 as described earlier.21 The electrospun fibers were collected on the surface of an earthed bath of phosphate buffered saline (PBS) and wound onto plastic carriers. The carriers were stored in PBS solution; the fibers were not allowed to dry out at any time. XHR SEM. Extra high resolution scanning electron microscopy (XHR SEM) was performed utilizing the MagellanTM 400L. Samples were prepared by drying a piece of fiber on filter paper followed by vacuum gold plating. Staining Methods. (1) Methylene Blue Staining: A stock solution of 0.1% (w/v) methylene blue and 2% (w/v) sodium citrate was added to an equal volume of cellular suspension, incubated for 10 min, washed and examined under light microscopy. Viable cells were unstained and dead cells were stained blue.22 B

DOI: 10.1021/acs.biomac.5b00970 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

Figure 2. Yeasts budding within electrospun microfibers containing C. tropicalis cells: (A) light microscope image taken immediately after the process; (B) 3 weeks post process.

■

(2) CTC viability stain: An amount of ∼5 μL 50 mM CTC (5cyano′-2,3-di(p-tolyl)tetrazolium chloride) (Sigma-Aldrich #94498) was sufficient to cover the yeast-containing tubes and was applied directly to a piece of the fiber on a glass slide. After incubation for 1.5 h at room temperature in the dark, the yeast-containing tubes were visualized by confocal microscopy. At least 10 fields were used for viability percent estimation. (3) LIVE/DEAD BacLight Bacterial Viability dye (Life technologies) was used to determine membrane integrity. Presto Blue Viability Assay. Presto blue (Invitrogen #A13261) was added to a final concentration of 10% directly to the yeast culture in 96-well plate. End point fluorescence was measured at time points for 2 h every 20 min with a BMG fluostar fluorimeter with an excitation wavelength of 544 nm and emission 590 nm. For the toxicity test, E. coli cells (from a laboratory collection) were grown to stationary phase. Equal amounts were taken for each well, centrifuged, and resuspended in minimal medium (as mentioned above); and appropriate concentrations of OMW were added. The test was done in triplicates. Phenol Determination. Phenol concentration was measured chemically by condensation of phenol with 4-aminoantipyrene followed by oxidation with alkaline potassium ferricyanide.23 Glycine buffer was adjusted to pH 9.7, and the measurement accuracy was ±0.1 mg/L. The optical density was measured at 505 nm in a Shimadzu UV mini 1240 Spectrophotometer. Phenol determination using HPLC was based on Comanet et al.24 Experiments with immobilized cells were done by immersing plastic carriers wrapped with the fibers in 10 mL sterile minimal medium in a 50 mL tube; phenol was added to a final concentration of 30−300 mg/L. For planktonic cells, a 24 h inoculum was centrifuged and resuspended with fresh phenol containing medium. Samples were taken periodically, treated with TCA, and frozen until tested. HPLC Analysis of OMW. Industrial OMW was obtained from a local medium-scale olive oil factory in central Israel. The sample was autoclaved and frozen. HPLC analysis was done according to Obied et al.25 Batch Fermentation. Studies of ethanol production were carried out at anoxic conditions (50 mL tubes containing 50 mL of CASO medium) with 10% glucose (w/v). The inoculum was prepared from an overnight preculture of yeast. After short initial shaking, the tubes were placed in an incubator at 37 °C for 48 h and sampled. All functions were measured comparing immobilized and planktonic cells. The cell count was normalized per 100 μL cell pellet and was estimated as ∼108 cells per mL. Ethanol Assay. Ethanol production was assayed with Megazyme International (Ireland) Ethanol Assay Kit (K-ETOH 1401) according to the manufacturer’s manual. Samples were taken directly from the liquid culture, centrifuged, and diluted to reach values in the linear range.

RESULTS AND DISCUSSION

3.1. Microtube Morphology and Cell Distribution within the Microtubes. Core−shell fibers were fabricated with the polymeric solutions described in the methods section. Fibers were uniform with an average diameter of 10 μm (Figure 1A). The fibers had porous walls of nano scale size due to PEG added to the shell solution, and the estimated pore sizes averaged 40 nm (Figure 1B). The pores allow for transfer of solubles but hinder cell leakage.26 The core PVP solution contained 20 vol % yeast cells. The cells within the fibers were evenly distributed, and no cells could be found exterior to the tubes (Figures 1C and 2A). After 3 days of incubation in rich medium, budding and division of the yeast cells were observed inside the tubes (Figure 2B). This indicates that the cells divide, and the internal space of the tubes is filled with cells post electrospinning. With longer storage periods (several months), the fibers maintain high cell density. The encapsulation method used in this study was chosen due to several factors: (1) The most striking result was the fact that the cells not only survived the process of electrospinning, but they multiplied, thus illustrating the first advantage of coelectrospun yeast. (2) The use of an insoluble polymer for the encapsulation, which provides the opportunity for water-based applications, as opposed to alginate/carrageenan/PVA and other polymers that are widely used today. (3) Due to the water-soluble PVP in the core, it is possible for the PVP to diffuse out through the shell wall, so with time there is no hindrance of water movement in the microtubes (data not shown). (4) A relatively high cell-to-encapsulating polymer ratio of 1:2.5 (by volume) is obtained. 3.2. Viability of Encapsulated Cells. Determination of cell viability imposed difficulties for several reasons: (a) encapsulated cells are difficult to quantify directly, because colony forming units on plates or slide counting of cells is not possible since the outer membrane is made of an insoluble polymer. (b) The polymers used interfere with conventional microscopic methods due to chemical dye absorbance of the outer membrane polymer and also autofluorescence at many wavelengths (green-red spectrum) and a complex and interlacing 3D structure. (c) Since there was limited certainty of viability assays based on a single enzymatic activity, several complementary methods were used. Viability was tested by examining respiration, enzymatic activity, and membrane integrity. The CTC is a tetrazolium salt reduced to formazan by the bacterial electron transport system. Direct microscopic C

DOI: 10.1021/acs.biomac.5b00970 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

Figure 3. Assessment of electrospun cell viability. (A) Control CTC stain of cells taken from an agar plate. (B) 3-week-old electrospun cells with CTC stain. (C) Control yeast cells - methylene blue staining. (D) Electrospun cells with methylene blue.

Figure 4. PI stained cells: (A) fluorescent image; (B) bright field overlay.

being used for their synthetic activity. Since the method requires that the cells withstand a high voltage electric field, the main initial question is the viability of the organisms after the electrospinning process. With yeast cells, it is clear that they survive the process as seen by their subsequent budding. However, other studies have indicated that restrictive environments arrest yeast cell cycles due to defects in bud emergence or actin perturbation.28 Furthermore, Nagarajan et al., who investigated the senescence of Saccharomyces cerevisiae when encapsulated in alginate, found that these cells in a system in which they did not divide but were not starved maintained their viability at a 95% level for 17 days, whereas free cells in suspension were less than 10% viable. Interestingly, their pattern of gene expression was different from that of starved or growing cells.29 In our study, budding, metabolic viability, as well as membrane integrity were observed. Cell cycle was not arrested, and all assayed viability parameters remained high

quantification showed that 95% of the cells are actively respiring (Figure 3A,B). Methylene blue is metabolized only in live cells, allowing for a simple measure of viability/survivalrate of C. tropicalis. It was estimated that about 90% of the cells were alive (destained) and thus have similar metabolic viability as when free cells are taken from an agar plate (Figure 3C,D). For membrane integrity - the high autofluorescence of the shell polymer in the green color range disturbed the visualization provided by the LIVE/DEAD viability kit SYTO 9 component. The second component (propidium iodide, PI) was visible (Figure 4), The PI showed staining, which suggests that some cells have compromised membranes. However, it has been proposed in previous studies that PI may stain components of intact yeast cell wall or membrane.27 In addition, many PI unstained cells may be seen. Although bacteria have been featured as agents of bioremediation, eukaryotic microorganisms, yeast, and algae can provide novel solutions to both bioremediation as well as D

DOI: 10.1021/acs.biomac.5b00970 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

with OMW showed substantial degradation of polyphenols. HPLC analysis of OMW incubated with C. tropicalis indicated degradation of several phenolic compounds (Figure 6A,B). Furthermore, the treatment of OMW by electrospun C. tropicalis completely reduced toxicity toward E. coli (Figure 6C). Since an important aspect of OMW pollution is its toxicity toward natural beneficial organisms, the ability to reduce this ecological effect is of significance. Ethanol Production. Ethanol production was tested in electrospun C. tropicalis and S. cerevisiae. For both yeasts, the electrospun immobilized cells produced less ethanol for the same incubation time (Table 1).

throughout 3 months of storage (the composite was stored in 4 °C in a sterile 1xPBS solution). 3.3. Immobilized Cell Functions. The main emphasis in earlier bacterial studies has been to look at the utilization of electrospun bacteria for bioremediation. In this study, several of the yeast’s activities that degrade pollutants were tested on pure phenol and phenolic compounds found in OMW. Phenol Biodegradation . The capability of C. tropicalis to degrade phenol is well established.30 In this study, planktonic cells biodegraded all the phenol in a solution within 24 h. However, electrospun cells degraded 45% of the added phenol after 48 h and 60% after 72 h of incubation (Figure 5). The electrospun cells thus degraded phenol with lower efficiency than free cells.

Table 1. Comparison of Suspended and Electrospun Cell Activitya function ethanol production

phenol degradation

planktonic C. tropicalis: 14 ± 0.1 [g/L] S. cerevisiae: 16 ± 0.1 [g/L] 100% after 48 hb

electrospun C. tropicalis: 6 ± 0.1 [g/L] S. cerevisiae: 3.5 ± 0.1 [g/L] 40% after 48 hb

a Compared by estimated cell quanity. bDrop in concentration as compared to initial.

Although electrospun yeast cells are viable, they have lower efficiency in all the activities tested. One reason for this phenomenon could be the challenge to accurately quantify the activity per cell. Although the cell number was estimated in our study, the fact that the cells multiply inside the polymer in subsequent storage prevents exact quantification. One question is whether the unique environment of the encapsulated cells could inhibit the dispersion of products yielding feedback inhibition of the enzymes involved.32 Another possibility is that the intricate structure of polymer mats could contain trapped air or liquid pockets that may inhibit fermentation. On the other hand, we believe that the emphasis on quantitative activity may be less important than qualitative aspects in the application of electrospun yeast.

Figure 5. Phenol biodegradation by planktonic and immobilized C. tropicalis; initial concentration: 100 mg/L in CASO medium.

OMW Biodegradation. OMW is an industrial effluent high in natural polyphenols. This effluent presents a serious environmental pollution problem, as its biological and chemical oxygen demand (COD) are high 100 and 220 mg O2 L−1 respectively.31 Alginate immobilized C. tropicalis was shown previously to biodegrade phenols and reduce COD and toxicity to microorganisms of OMW under metabolic induction.12 In this work, incubation of electrospun immobilized C. tropicalis

Figure 6. Biodegradation of wastewater phenols. (A) HPLC analysis of OMW components. (B) HPLC analysis after incubation with C. tropicalis; arrows point at changed peaks. (C) Toxicity evaluation of treated OMW by measure of E. coli viability. E

DOI: 10.1021/acs.biomac.5b00970 Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

■

Candida sp. has been noted to synthesize xylitol, fatty acids, long chain hydrocarbons, terpenes, and multiple precursors for the pharmaceutical and plastic industry, and in one case, even nanoparticles.16,33−35 Huf et al.,36 have reviewed the biotechnological synthesis of dicarboxylic acids (DCA) since these compounds find usage in a great number of polymeric formulations, especially as “green polymers”. In this review, C. tropicalis plays a central role, featuring the basic genetic manipulations to both cancel out alkane degradation and allow accumulation of DCAs for the production of industrial quantities of these compounds. Even though the biological production generally results in far fewer side products, purification still remains energy and labor consuming. Electrospun yeast will probably not be competitive with such production levels, but since many of the synthetic studies have been carried out in bioreactors of various configurations, it is possible that electrospun yeast may add the advantages of spatial separation. Electrospun yeast may also contribute to the solution of the problem of postproduction purification. This may be through the development of “smart polymers,” which may open up the fiber encapsulated yeast to substrate and release specific products in a controlled fashion. Previously live bacterial cells were encapsulated within polymer fibers in two forms: (A) one polymer type−monolithic fibers and (B) two-polymer fibers (core−shell). The first method requires a water-soluble polymer (PEO, PVA, silica), thus its first disadvantage is due to the liquid environment required for the application of such composites due to the inevitable breakdown of polymer structure. In order to overcome this challenge, several studies added a cross-linking step, which requires harsh organic solvents such as acetone or glutaraldehyde.7,37 Although some microorganisms could survive these processes in a fair percentage of viable cells, their lifespan and activity were limited.37 The second method (core−shell) allows the creation of hydrophilic core (PVA/ PVP) and a hydrophobic porous shell (PEG/PVDF) in one step. This method allows a fairly easy and nontoxic way to encapsulate cells within insoluble polymer fibers. We have shown that microbial cells remain viable and active in such environments.8 Major challenges for future work are the study of microenvironmental biology of these encapsulated yeast and its effect on their overall metabolism, synthetic capacity, gene activity, and if and how biofilm is formed. Novel polymeric formulations might enable the retention of activities similar to cells in suspension. Another interesting possibility is the integration of natural and synthetic polymers.

Article

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. *E-mail: [email protected]. Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We thank Diana Ickowicz for the OMW HPLC. We wish to acknowledge the advice and assistance of Professors Oded Shoseyov, Ariel Kushmaro, and Ophry Pines. We would like to thank Roi Ramot for the 3D illustration. This research was funded by the GIP program of the Deutsche Forschungsgemeinschaft (DFG) German Research Foundation.

■ ■

ABBREVIATIONS OMW, Olive mill wastewater REFERENCES

(1) Cassidy, M.; Lee, H.; Trevors, J. T. Environmental applications of immobilized microbial cells: a review. J. Ind. Microbiol. 1996, 16, 79− 101. (2) Adam, W.; Lukacs, Z.; Harmsen, D.; Saha-Moller, C. R.; Schreier, P. Biocatalytic asymmetric hydroxylation of hydrocarbons with the topsoil-microorganism Bacillus megaterium. J. Org. Chem. 2000, 65, 878−82. (3) Dervakos, G. A.; Webb, C. On the merits of viable-cell immobilisation. Biotechnol. Adv. 1991, 9, 559−612. (4) Gadkari, D. Nitrification in the presence of soil particles, sand, alginate beads and agar strands. Soil Biol. Biochem. 1990, 22, 17−21. (5) Keweloh, H.; Heipieper, H.-J.; Rehm, H.-J. Protection of bacteria against toxicity of phenol by immobilization in calcium alginate. Appl. Microbiol. Biotechnol. 1989, 31, 383−389. (6) Knierim, C.; Greenblatt, C. L.; Agarwal, S.; Greiner, A. Blocked Bacteria Escape by ATRP Grafting of a PMMA Shell on PVA Microparticles. Macromol. Biosci. 2014, 14, 537−545. (7) Canbolat, M. F.; Gera, N.; Tang, C.; Monian, B.; Rao, B. M.; Pourdeyhimi, B.; Khan, S. A. Preservation of Cell Viability and Protein Conformation on Immobilization within Nanofibers via Electrospinning Functionalized Yeast. ACS Appl. Mater. Interfaces 2013, 5, 9349−9354. (8) Klein, S.; Kuhn, J.; Avrahami, R.; Tarre, S.; Beliavski, M.; Green, M.; Zussman, E. Encapsulation of bacterial cells in electrospun microtubes. Biomacromolecules 2009, 10, 1751−6. (9) Gensheimer, M. B.; Becker, M.; Brandis-Heep, A.; Wendorff, J. H.; Thauer, R. K.; Greiner, A. Novel Biohybrid Materials by Electrospinning: Nanofibers of Poly(ethylene oxide) and Living Bacteria. Adv. Mater. 2007, 19, 2480−2482. (10) Dror, Y.; Salalha, W.; Avrahami, R.; Zussman, E.; Yarin, A. L.; Dersch, R.; Greiner, A.; Wendorff, J. H. One-step production of polymeric microtubes by co-electrospinning. Small 2007, 3, 1064−73. (11) Greiner, A.; Wendorff, J. H. Electrospinning: A fascinating method for the preparation of ultrathin fibres. Angew. Chem., Int. Ed. 2007, 46, 5670−5703. (12) Ettayebi, K.; Errachidi, F.; Jamai, L.; Tahri-Jouti, M. A.; Sendide, K.; Ettayebi, M. Biodegradation of polyphenols with immobilized Candida tropicalis under metabolic induction. FEMS Microbiol. Lett. 2003, 223, 215−9. (13) Jiang, Y.; Wen, J.; Lan, L.; Hu, Z. Biodegradation of phenol and 4-chlorophenol by the yeast Candida tropicalis. Biodegradation 2007, 18, 719−29.

■

CONCLUSIONS In the current study we report the assembly of a novel yeastpolymer composite and demonstrate its usage in biotechnological activities. The electrospun composite has a unique structure that allows for great surface area and free liquid transfer, but hinders cell leakage. In addition, it provides the yeast a PVP core that enables them to reside in an aqueous environment. As a result, we could detect extensive yeast budding in the composite and cellular function in the biodegradation of phenols and production of ethanol. Moreover, with the yeast’s modification of olive water waste, it became less toxic for E. coli. These findings demonstrate that the coelectrospun composite can be a promising platform for creation of active microbe−polymeric systems that can be used in water-based applications. F

DOI: 10.1021/acs.biomac.5b00970 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

astaxanthin in the food yeast Candida utilis. Appl. Environ. Microbiol. 1998, 64, 1226−9. (35) Hubenova, Y.; Mitov, M. Potential application of Candida melibiosica in biofuel cells. Bioelectrochemistry 2010, 78, 57−61. (36) Huf, S.; Krugener, S.; Hirth, T.; Rupp, S.; Zibek, S. Biotechnological synthesis of long-chain dicarboxylic acids as building blocks for polymers. Eur. J. Lipid Sci. Technol. 2011, 113, 548−561. (37) Gensheimer, M.; Brandis-Heep, A.; Agarwal, S.; Thauer, R. K.; Greiner, A. Polymer/Bacteria Composite Nanofiber Nonwovens by Electrospinning of Living Bacteria Protected by Hydrogel Microparticles. Macromol. Biosci. 2011, 11, 333−337.

(14) Farag, S.; Soliman, N. A. Biodegradation of crude petroleum oil and environmental pollutants by Candida tropicalis strain. Braz. Arch. Biol. Technol. 2011, 54, 821−830. (15) Rehman, A.; Farooq, H.; Shakoori, A. R. Copper Tolerant Yeast, Candida tropicalis, Isolated from Industrial Effluents: its Potential use in Wastewater Treatment. Pakistan J. Zool. 2007, 39, 405−412. (16) Dasilva, S. S.; Afschar, A. S. Microbial-Production of Xylitol from D-Xylose Using Candida-Tropicalis. Bioprocess Eng. 1994, 11, 129−134. (17) PROSODOL Website. Olive Oil Production in the Mediterranean. http://www.prosodol.gr/?q=node/203. (18) Rouvalis, A.; Iliopoulou-Georgudaki, J. Comparative assessment of olive oil mill effluents from three-phase and two-phase systems, treated for hydrogen production. Bull. Environ. Contam. Toxicol. 2010, 85, 432−6. (19) International Programme on Chemical Safety Environmental Health Criteria 161, Phenol; World Health Organization: Geneva, Switzerland, 1994. (20) Asfi, M.; Ouzounidou, G.; Moustakas, M. Evaluation of olive oil mill wastewater toxicity on spinach. Environ. Sci. Pollut. Res. 2012, 19, 2363−2371. (21) Nardi, A.; Avrahami, R.; Zussman, E.; Rokem, J. S.; Greenblatt, C. L. Phenol Biodegradation by Corynebacterium glutamicum Encapsulated in Electrospun Fibers. J. Environ. Prot. 2012, 3, 164−168. (22) Lange, H.; Bavouzet, J. M.; Taillandier, P.; Delorme, C. Systematic error and comparison of four methods for assessing the viability of Saccharomyces cerevlsiae suspensions. Biotechnol. Tech. 1993, 7, 223−228. (23) Der Yang, R.; Humphrey, A. E. Dynamic and steady state studies of phenol biodegradation in pure and mixed cultures. Biotechnol. Bioeng. 1975, 17, 1211−1235. (24) Coman, V.; Moldovan, Z. RP-HPLC method for the separation of some phenol derivatives using gradient elution and UV detection. J. High Resolut. Chromatogr. 2000, 23, 699−701. (25) Obied, H. K.; Allen, M. S.; Bedgood, D. R.; Prenzler, P. D.; Robards, K.; Stockmann, R. Bioactivity and analysis of biophenols recovered from olive mill waste. J. Agric. Food Chem. 2005, 53, 823− 37. (26) Dror, Y.; Kuhn, J.; Avrahami, R.; Zussman, E. Encapsulation of enzymes in biodegradable tubular structures. Macromolecules 2008, 41, 4187−4192. (27) Corliss, D. A.; White, W. E., Jr. Fluorescence of yeast vitally stained with ethidium bromide and propidium iodide. J. Histochem. Cytochem. 1981, 29, 45−8. (28) Suzuki, M.; Asada, Y.; Watanabe, D.; Ohya, Y. Cell shape and growth of budding yeast cells in restrictive microenvironments. Yeast 2004, 21, 983−9. (29) Nagarajan, S.; Kruckeberg, A. L.; Schmidt, K. H.; Kroll, E.; Hamilton, M.; McInnerney, K.; Summers, R.; Taylor, T.; Rosenzweig, F. Uncoupling reproduction from metabolism extends chronological lifespan in yeast. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, E1538−47. (30) Krug, M.; Ziegler, H.; Straube, G. Degradation of PhenolicCompounds by the Yeast Candida-Tropicalis Hp-15 0.1. Physiology of Growth and Substrate Utilization. J. Basic Microbiol. 1985, 25, 103− 110. (31) Paixao, S. M.; Mendonca, E.; Picado, A.; Anselmo, A. M. Acute toxicity evaluation of olive oil mill wastewaters: A comparative study of three aquatic organisms. Environ. Toxicol. 1999, 14, 263−269. (32) Amenaghawon, N. A.; Okieimen, C. O.; Ogbeide, S. E. Kinetic Modelling of Ethanol Inhibition during Alcohol fermentation of Corn Stover using Saccharomyces Cerevisiae. Int. J. Eng. Res. Ind. Appl. 2012, 2 (4), 798−803. (33) Picataggio, S.; Rohrer, T.; Deanda, K.; Lanning, D.; Reynolds, R.; Mielenz, J.; Eirich, L. D. Metabolic engineering of Candida tropicalis for the production of long-chain dicarboxylic acids. Bio/ Technology 1992, 10, 894−8. (34) Miura, Y.; Kondo, K.; Saito, T.; Shimada, H.; Fraser, P. D.; Misawa, N. Production of the carotenoids lycopene, beta-carotene, and G

DOI: 10.1021/acs.biomac.5b00970 Biomacromolecules XXXX, XXX, XXX−XXX