Nitrate removal from liquid effluents using microalgae

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Nitrate removal from liquid effluents using microalgae immobilized on chitosan nanofiber mats Ela Eroglu,a,b Vipul Agarwal,a Michael Bradshaw,a Xianjue Chen,a Steven M. Smith,*b Colin L. Raston*a and K. Swaminathan Iyera Received 25th June 2012, Accepted 8th August 2012 DOI: 10.1039/c2gc35970g

Mats of electrospun chitosan nanofibers were found effective in immobilizing microalgal cells. These immobilized microalgal cells were also used as a durable model system for wastewater treatment which has been demonstrated by the removal of around 87% nitrate from liquid effluents ([NO3−N]initial 30 mg L−1). Virtuous nitrate removal rates were derived by superimposing physicochemical adsorption and biological nutrient consumption phenomena for chitosan and microalgae, respectively. Wastewater treatment focuses on eliminating unwanted chemicals and/or biological impurities from contaminated water. In general the treatment methods are mainly based on the separation of pollutants from the wastewater with a requirement for a further processing stage to eliminate these pollutants.1 Integrated wastewater treatment processes are important in eliminating undesired species, ideally converting them into valuable products. As relatively recent bioprocesses, algal cultivation in wastewaters has a combination of several advantages such as wastewater treatment and simultaneous algal biomass production, which can be further exploited for biofuel production, food additives, fertilizers, cosmetics, pharmaceuticals, and other valuable chemicals.2 However, there are inherent difficulties associated with algae-based bioprocessing in the harvesting, dewatering and processing of the algal biomass. Immobilization of the cells on solid surfaces confer advantages over free cells in suspension, namely the immobilized cellular matter occupy less space, require smaller volume of growth medium, are easier to handle, and can be used repeatedly for product generation.3,4 Moreover, immobilization can also increase the resistance of cell cultures to harsh environmental conditions such as salinity, metal toxicity and variations in pH.3,4 Entrapment is one of the most common immobilization methods which involves capturing the cells in a three dimensional gel matrix, made of polymeric materials or inorganic spheres.4 Both synthetic polymers (e.g., acrylamide, a

Centre for Strategic Nano-Fabrication, School of Chemistry and Biochemistry, The University of Western Australia, M313, 35 Stirling Highway, Crawley, WA 6009, Australia. E-mail: [email protected]; Tel: +61 8 6488 3045; Fax: +61 8 6488 8683 b ARC Centre of Excellence in Plant Energy Biology, The University of Western Australia M313, 35 Stirling Highway, Crawley, WA 6009, Australia. E-mail: [email protected]; Tel: +61 8 6488 4403; Fax: +61 8 6488 4401 2682 | Green Chem., 2012, 14, 2682–2685

polyurethane, polyvinyl) and natural polymers (e.g., collagen, agar, cellulose, alginate, carrageenan) have been used for this purpose.5 Several studies have been reported on wastewater treatment involving the entrapment of microalgae cultures inside alginate beads, porous glass, and several synthetic polymers.6–8 However, most attempts to immobilize viable algae cells inside such insoluble materials have limitations, with the encapsulating materials having volume/surface ratios usually orders of magnitude larger than thin films. As a consequence, algal viability is mostly reported to decrease which relates to the need for the nutrients or reactants to diffuse far into the material to reach the algal cells.5 We have developed a new technique to overcome these problems using electrospun nanofiber mats as the matrix for immobilizing the algal cells, with an overall strategy to combine wastewater treatment processing with algal harvesting in a single process. Electrospun nanofibers of chitosan were employed as a polymer/matrix support for green microalgae in the current study. Chitosan is composed of D-glucosamine and N-acetyl-Dglucosamine, and is formed by the deacetylation of chitin (β-Nacetyl-D-glucosamine polymer).9 Chitin is usually extracted from the exoskeletons of crustaceans (e.g., crab, lobster and shrimp) and even from the cell walls of fungi.9,10 Chitosan is non-toxic and biodegradable, and can be used as an animal feed.9 Furthermore, it can be used as a coagulant for wastewater treatment and for the recovery of waste sludge.11 The removal of nitrate ions is regulated by law which relates to its hazardous effects on human health and the environment. Several methods have been reported for the removal of nitrate from water bodies, including biological denitrification,12 chemical reduction,13 electrodialysis,14 and a combined bioelectrochemical/adsorption process.15 In this study, we aimed to superimpose the treatment efficiencies of microalgae in reducing nitrate and electrostatic binding of the nitrate ion by chitosan, i.e. combining biological and chemical processing. Chlorella vulgaris cultures were used as the green microalgae which were originally obtained from the Australian National Algae Culture Collection at CSIRO, Tasmania. Cell growth was carried out under artificial diurnal-illumination (16 h light/8 h dark cycle) at around 22 °C. Electrospun chitosan mats were fabricated following the procedure optimized in previous studies.16,17 This involved dissolving chitosan powder (6 wt%) in a mixture of trifluoroacetic acid (TFA) and dichloromethane (DCM) (70 : 30 v/v), with 5.4% (v/v) addition of glutaraldehyde This journal is © The Royal Society of Chemistry 2012

solution (25% in H2O) immediately prior to electrospinning.17 This was deemed necessary to cross-link the chitosan to avoid polymer breakup and dissolution. The experimental settings of electrospinning processes were as follows: (i) syringe pump speed: 0.1 mm min−1, (ii) voltage: 18 kV, (iii) distance between the target and the tip of the syringe: 11 cm, (iv) target speed: 1 m min−1, (v) traverse speed: 0.5 cm min−1. Fiber mats were annealed overnight to remove any remaining solvent and stored until required. After electrospinning, 2 mL of algae solution in its exponential phase of growth with a total chlorophyll content (Chl a and b) of ∼2 mg L−1, was placed onto a cut out rectangular chitosan-mat (3 × 2 cm) and left at room temperature for approximately 48 hours to allow sufficient attachment of algae cells to the surface of the mat. Microalgae cell walls contain various polysaccharides, which are compatible with the surface of the chitosan nanofibers.18,19 The presence of negative surface charge on the surface of Chlorella cells, arising from dissociation of uronic acid groups, and/or the presence of sulfate groups for example,18 provide electrostatic attraction to the positively charged primary amine groups of chitosan not involved in the above cross linking. Moreover, the negatively charged surface of the microalgae can also result in binding metal ions, thereby providing an opportunity for biosorption applications, along with the removal of nitrate ions as functional algal cells.18–20 This bionano-composite material was then placed into nitrate containing artificial growth medium which contained mainly phosphates, nitrates, carbonate buffer, micronutrients and vitamins,21 with an initial nitrate-nitrogen concentration of around 30 mg L−1. Nitrate-nitrogen (NO3−-N) term refers to the amount of nitrogen (N) in liquid solutions coming from nitrate ions (NO3−). This nitrate-nitrogen concentration is within the range of other algal nitrate removal studies.2,22 Furthermore, it simulates the range of nitrate content present in ground-waters (∼0.1–50 mg L−1 NO3−-N)23 and sewage treatment plant effluents. The regulatory limit for the maximum contaminant levels of [NO3−-N] in public drinking water is 10 mg L−1, as established by the United States Environmental Protection Agency (EPA).24,25 During the algal growth, the pH of the medium was kept around 6.5–7.0 by the addition of dilute hydrochloric acid (HCl) when necessary. The colorimetric “cadmium reduction method” was employed for the nitrate-nitrogen analysis, using chemicalkits in the form of powder pillows (HACH®, NitraVer Nitrate Reagent) and a colorimeter (HACH® DR/870).26 For comparative purposes, a control experiment with a chitosan nanofiber mat devoid of algae culture was also treated under the same conditions. Scanning electron microscopy (SEM) analyses were acquired using a Zeiss 1555 VP-FESEM, while the accelerating voltage was changed between 3 to 5 kV. The air-dried samples were coated with approximately 3 nm layer of platinum before imaging. A NanoMan AFM system (Veeco Instruments Inc.) was used for the atomic force microscopy (AFM) analysis, operating under the tapping mode. Chlorophyll content of the cells was analyzed using spectrophotometric measurements of methanol extracts obtained from the algal culture pellets.27 A challenge in the present work was to fabricate an insoluble, fibrous structure with sufficient porosity, which can facilitate the diffusion of nutrients and cellular products between the environment and the algae. Fig. 1a and b show scanning electron This journal is © The Royal Society of Chemistry 2012

Fig. 1 SEM images of as prepared electrospun chitosan nanofibers at low and high magnifications with scale bars of (a) 1 μm, and (b) 10 μm, respectively.

Fig. 2 (a) SEM images of porous and swollen chitosan nanofiber mats, after exposure to nitrate containing media for two days, (b) SEM images of chitosan nanofibers surrounding individual, and (c) multiple algae cells. Scale bars are given as 1 μm.

microscopic (SEM) images of the nanofiber structure of the chitosan mats after the electrospinning process. The diameter of the fibers was between 50 to 180 nm, with an average diameter of around 91 nm. After placing the nanofiber mat into aqueous solution, the fibers gradually swelled with a significant increase in porosity of the material, Fig. 2a, and became an effective support matrix for the C. vulgaris cells, which have the expected diameter from SEM images around 3–4 μm, Fig. 2b and c. This porous structure has an advantage for facilitating the diffusion of materials such as nutrients and waste products between the environment and the algae, while the replication of algal cells is accomplished on the surface of the nanofiber mat, Fig. 2c. The height profile measurements obtained by AFM analyses established the thickness of the chitosan mat at close to 400 nm, Fig. 3a, with the overall thickness for the algae attached chitosan mat at 4.3 μm, Fig. 3b. The difference in height is consistent with the attachment of a single layer of individual C. vulgaris cells on the nanofiber mats. Fig. 4 shows the optical images of algae cells attached to the surface of chitosan mats (a) initially, (b) after 3 days, and (c) after 10 days from the start of the growth experiments in 40 mL liquid media. Note the increasing green color on the surface is due to the increased concentration of algal cells on the chitosan mat with respect to time. Detailed imaging is given as SEM images in Fig. 5. The amount of algae cells on a chitosan mat with same dimensions yielded around 4 cells per 100 μm2 for a 3 day old sample, Fig. 5c, whereas this increased to around Green Chem., 2012, 14, 2682–2685 | 2683

Fig. 3 AFM topographic mapping of chitosan nanofiber mats (4 × 4 μm) without, (a) and with, (b) algal cells.

Fig. 6 Nitrate-nitrogen (NO3−-N) concentration (mg L−1) of algal medium versus time. Chitosan nanofibers devoid of algal cells are represented with triangles, whereas those with immobilized algal cells are shown as rectangles. Fig. 4 Progress of the algal growth on the surface of chitosan mats: (a) initially, (b) after 3 days, (c) after 10 days of the growth experiment.

Fig. 5 SEM images of immobilized C. vulgaris cells on the surface of chitosan nanofiber mats after different time intervals. (a and c) are for mats after 3 days, and (b and d) are for mats 10 days old.

20 cells per 100 μm2 by the 10th day of the treatment process, Fig. 5d. Fig. 6 shows the nitrate-nitrogen concentration versus time in the absence of algae (triangles) or algae attached (rectangles) nanofiber mats. After the insertion of this bionano-composite into the liquid media (Vtotal: 40 mL), around 30% of the initial nitrate value was decreased within the first 2 days. This reduction in nitrate is mainly caused through the uptake by the chitosan nanofibers rather than the algal cultures, as a physicochemical adsorption process. A similar pattern was also observed for the mats devoid of algal cells where after the second day there was no further nitrate removal. In contrast the algae containing mats continued their nitrate uptake, being used in their cellular metabolism for replication, building more biomass and energy products. 2684 | Green Chem., 2012, 14, 2682–2685

Amino groups in chitosan are protonated at acidic to neutral pH conditions,28 which enhance the adhesive properties of chitosan by increasing its tendency to attach negatively charged entities which in this case are algal cell walls and nitrates. Chlorella vulgaris cell walls are known to be highly negatively charged with a zeta potential of around −30 mV in neutral water.29 On the other hand, the zeta potential of the positively charged chitosan nanofibers is +20 mV at neutral pH. Matsumoto et al.30 reported the zeta potential values of chitosan nanofibers to be highly dependent on the pH of the media, with it increasing to +30 mV in pH around 5–6, whereas it drops to zero for pH values above 8.30 Algal growth tends to alkalify its medium, as the cellular uptake of anions (such as nitrates, phosphates, carbonates, etc.) is stabilized with equivalent amounts of hydroxyl (OH−) anion efflux.31 For this reason, we maintained the pH of the culture around 6.5–7 by the regular addition of dilute HCl during the current study. Due to the nature of HCl, several other acidifying agents (such as CO2) can be considered for any future developments and advanced scale-up processes for municipal and/or industrial wastewater samples. Clearly the presence of the nanofiber mat in the liquid environment is responsible for the initial removal of nitrate while the continued growth of algae subsequently consumes the remaining nitrate in further stages with a slower rate. Overall nitrate removal rates were calculated as 32 ± 3%, and 87 ± 4%, for the “microalgae-absent” and “microalgae-attached” chitosan mats, respectively. Several studies have already been reported on wastewater treatment with immobilized microorganisms. Fierro et al.22 investigated the effect of nitrate removal by Scenedesmus spp. cyanobacterial cells immobilized within spherical chitosan beads. They achieved 70% nitrate removal for the immobilized cultures, while 20% of the initial nitrate content was removed by the blank chitosan beads. In another study, Mallick and Rai32 also achieved relatively higher nitrate removal rates (73%) by Anabaena doliolum and Chlorella vulgaris cells immobilized in chitosan beads compared to the cells immobilized in other types of gels made of alginate, carrageenan or agar. De-Bashan et al.33 achieved only 15% nitrate removal for co-immobilized This journal is © The Royal Society of Chemistry 2012

microorganisms (Chlorella vulgaris with a growth-promoting bacterium Azospirillum brasilense) within alginate beads. At the other end of the scale, Tam and Yong34 reported complete nitrate removal using immobilized C. vulgaris cells within calcium alginate beads. The treatment efficiency of our current method is comparable with that of these aforementioned methods, although large variations among the experimental parameters; including wastewater composition, microbial species, duration of the process, type of bioreactor, chemical composition and shape of the immobilization matrix, make direct comparison difficult. In summary, we have established the use of cross-linked chitosan nanofiber mat as a water-insoluble and non-toxic support for algal growth and nitrate removal from waters. Algal growth on a support material can lead to combine algal harvesting, dewatering, and processing steps in a single stage. This bionano-composite material is potentially an attractive, simple and highly durable polymer, with the mats still retaining their integrity after six months in contact with an aqueous solution, and has promise for industrial and/or municipal wastewater treatment processes.

Acknowledgements This work has been supported by The University of Western Australia and the Australian Research Council. We would like to acknowledge the facilities of the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterization & Analysis, The University of Western Australia, which was funded by the University, State and Commonwealth Governments. AFM images were performed at Curtin University.

Notes and references 1 Metcalf and Eddy, Inc, Wastewater Engineering: Treatment and Reuse, McGraw-Hill, New York, 4th edn, 2003. 2 N. Mallick, BioMetals, 2002, 15, 377. 3 L. Hall-Stoodley, J. W. Costerton and P. Stoodley, Nat. Rev. Microbiol., 2004, 2(2), 95. 4 Y. Liu, M. H. Rafailovich, R. Malal, D. Cohn and D. Chidambaram, Proc. Natl. Acad. Sci. U. S. A., 2009, 106(34), 14201. 5 M. S. A. Hameed and O. H. Ebrahim, Int. J. Agri. Biol., 2007, 9, 183.

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6 K. Abe, E. Takahashi and M. Hirano, J. Appl. Phycol., 2008, 20, 283. 7 G. Schumacher and I. Sekoulov, Water Sci. Technol., 2002, 46, 83. 8 E. Delahaye, R. Boussahel, T. Petitgand, J. P. Duguet and A. Montiel, Desalination, 2005, 177, 273. 9 A. Lavoie and J. de La Noue, J. World Maricul. Soc., 1983, 14, 685. 10 A. Zamani, L. Edebo, B. Sjöström and M. J. Taherzadeh, Biomacromolecules, 2007, 8, 3786. 11 W. A. Bough, Process Biochem., 1976, 11, 13. 12 E. Wasik, J. Bohdziewcz and M. Blaszczyk, Process Biochem., 2001, 37, 57. 13 H.-Y. Hu, N. Goto and K. Fujie, Water Res., 2001, 35, 2789. 14 A. Elmidaoui, F. Elhannouni, S. M. A. Menkouchi Sahli, L. Chay, E. Elabbassi, M. Hafsi and D. Largeteau, Desalination, 2001, 136, 325. 15 Z. Feleke and Y. Sakakibara, Water Sci. Technol., 2001, 43, 25. 16 K. Ohkawa, D. I. Cha, H. Kim, A. Nishida and H. Yamamoto, Macromol. Rapid Commun., 2004, 25, 1600. 17 M. Bradshaw, J. Zou, L. Byrne, K. S. Iyer, S. G. Stewart and C. L. Raston, Chem. Commun., 2011, 47, 12292. 18 D. Kaplan, D. Christiaen and S. M. Arad, Appl. Environ. Microbiol., 1987, 53, 2953. 19 R. H. Crist, K. Oberholser, N. Shank and M. Nguyen, Environ. Sci. Technol., 1981, 15, 1212. 20 B. Volesky and Z. R. Holan, Biotechnol. Prog., 1995, 11, 235. 21 C. J. S. Bolch and S. I. Blackburn, J. Appl. Phycol., 1996, 8, 5. 22 S. Fierro, M. del P. Sanchez-Saavedra and C. Copalcua, Bioresour. Technol., 2008, 99, 1274. 23 U. N. Dwivedi, S. Mishra, P. Singh and R. D. Tripathi, in Environmental Bioremediation Technologies, ed. S. N. Singh and R. D. Tripathi, Springer, New York, 2007, ch. 16, pp. 353–389. 24 EPA, National Pesticide Survey: Project Summary, U.S. Environmental Protection Agency, Washington DC, 1990. 25 S. Ghafari, M. Hasan and M. K. Aroua, Bioresour. Technol., 2008, 99, 3965. 26 APHA, Standard Methods for the Examination of Water and Wastewater, American Public Health Association, Washington, DC, 18th edn, 1992. 27 H. K. Lichtenthaler and C. Buschmann, in Current Protocols in Food Analytical Chemistry, ed. R. E. Wrolstad, John Wiley & Sons Inc., New York, 2001, pp. F4.3.1–F4.3.8. 28 L.-Q. Wu, P. Gadre Anand, H. Yi, M. J. Kastantin, W. Rubloff Gary, E. Bentley William, F. Gregory Payne and R. Ghodssi, Langmuir, 2002, 18, 8620. 29 B.-M. Hsu, Parasitol. Res., 2006, 99, 357. 30 H. Matsumoto, H. Yako, M. Minagawa and A. Tanioka, J. Colloid Interface Sci., 2007, 310, 678. 31 J. Naus and A. Melis, Plant Cell Physiol., 1991, 32, 569. 32 N. Mallick and L. C. Rai, World J. Microbiol. Biotechnol., 1994, 10, 439. 33 L. E. de-Bashan, Y. Bashan, M. Moreno, V. K. Lebsky and J. J. Bustillos, Can. J. Microbiol., 2002, 48, 514. 34 N. F. Y. Tam and Y. S. Wong, Environ. Pollut., 2000, 107, 145.

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