- Email: [email protected]
Microalgae-associated bacteria play a key role in the flocculation of Chlorella vulgaris
Bioresource Technology 131 (2013) 195–201
Contents lists available at SciVerse ScienceDirect
Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Microalgae-associated bacteria play a key role in the flocculation of Chlorella vulgaris Jimin Lee a,b, Dae-Hyun Cho a, Rishiram Ramanan a, Byung-Hyuk Kim a, Hee-Mock Oh a,b, Hee-Sik Kim a,b,⇑ a b
Environmental Biotechnology Research Center, Korea Research Institute of Bioscience and Biotechnology, 125 Gwahak-ro, Yuseong-gu, Daejeon 305-806, Republic of Korea University of Science and Technology, 217 Gajeong-ro, Yuseong-gu, Daejeon, Republic of Korea
h i g h l i g h t s " Flocculation of axenic and xenic cultures of Chlorella vulgaris implicates bacteria. " DGGE analysis indicates presence of five species of microalgae-associated bacteria. " FACS treatment of xenic culture implicates three bacterial species in flocculation. " Bacteria and its extracellular substances increase floc size. " This study proves that bacteria play a major role in flocculation of microalgae.
a r t i c l e
i n f o
Article history: Received 16 July 2012 Received in revised form 22 October 2012 Accepted 28 November 2012 Available online 8 December 2012 Keywords: Algae Flocculation Chlorella vulgaris Microalgae-associated bacteria
a b s t r a c t Flocculation is most preferred method for harvesting microalgae, however, the role of bacteria in microalgal flocculation process is still unknown. This study proves that bacteria play a profound role in flocculating by increasing the floc size resulting in sedimentation of microalgae. A flocculating activity of 94% was achieved with xenic Chlorella vulgaris culture as compared to 2% achieved with axenic culture. Denaturing gradient gel electrophoresis (DGGE) analysis of 16S rRNA gene of xenic C. vulgaris culture revealed the presence of Flavobacterium sp., Terrimonas sp., Sphingobacterium sp., Rhizobium sp. and Hyphomonas sp. as microalgae-associated bacteria. However when Flavobacterium, Terrimonas, Sphingobacterium were eliminated by fluorescence activated cell sorter (FACS), flocculating activity reduced to 3%. Further studies with cell free extracts also suggest that bacterial extracellular substances might also have a role in enhancing flocculation. We conclude that the collective presence of certain bacteria is the determining factor in flocculation of C. vulgaris. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Microalgae are now receiving extensive global attention as a potential source for biofuel production after being touted as an alternative fuel source some decades ago (Benemann et al., 1977; Oswald and Golueke, 1960). The production process of biodiesel using microalgal biomass includes cultivation, harvest, oil extraction, and conversion. Harvesting of microalgae is one of the most critical steps which involves separation of biomass from culture medium and contributes about 20–30% of the total biomass production cost (Gudin and Thepenier, 1986; Uduman et al., 2010). Harvesting is especially critical when the product of interest is a relatively low-value product like biodiesel (Vandamme et al., ⇑ Corresponding author at: Environmental Biotechnology Research Center, Korea Research Institute of Bioscience and Biotechnology, 125 Gwahak-ro, Yuseong-gu, Daejeon 305-806, Republic of Korea. Tel.: +82 42 860 4326; fax: +82 42 879 8103. E-mail address: [email protected] (H.-S. Kim). 0960-8524/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2012.11.130
2010). The major challenges in harvesting microalgae is because of small size of microalgal single cell (typically a few micrometer) and its low concentration in the culture medium (0.5–2 g L1) (Schlesinger et al., 2012; Vandamme et al., 2011). Most common harvesting methods include flocculation, gravity sedimentation, centrifugation, filtration and ultrafiltration, sometimes with an additional flocculation step or with a combination of flocculation–flotation. Centrifugation is one of the preferred methods for microalgal cell recovery because of effectiveness and rapidness but it requires energy thereby increasing operating costs (Molina Grima et al., 2003). Membrane replacement is the major cost involved in filtration method and this also depends on the concentration of microalgae (Wilde et al., 1991). One of the most potential methods to reduce cost and energy usage during harvesting is flocculation (Wyatt et al., 2012). Algal cells carry negative charge on the cell surface preventing aggregation because of repulsion; however the addition of cationic metal ions such as Ca2+ and Fe3+ can neutralize this charge, leading to the aggregation of cells.
196
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
Various flocculation methods result in higher particle sizes to enable gravity sedimentation, centrifugal recovery, and filtration (Molina Grima et al., 2003). The harvesting of microalgal cells by flocculation is more convenient process than contemporary methods such as centrifugation and filtration because it allows the treatment of large quantities of microalgal culture (Lee et al., 1998) and can be applied to a wide range of species (Pushparaj et al., 1993). Among the various flocculants, Aluminium sulphate (Alum) is most widely used for removal of algae, because of ease of use and application (Ebeling et al., 2003; Schlesinger et al., 2012). However, it cannot be applied over a wide pH range, moreover, floc size with alum when compared to ferric flocs is smaller resulting in ineffective sedimentation (Ebeling et al., 2003). Other cations such as calcium and magnesium also have a positive effect on flocculation in high pH (Vandamme et al., 2012). In addition, cationic polymers such as chitosan (Divakaran and Sivasankara Pillai, 2002), or alkalis such as NaOH have been used to achieve better flocculation (Brennan and Owende, 2010). On the contrary, various species of algae have been reported to auto-flocculate (Spilling et al., 2011; Sukenik and Shelef, 1984). There have been reports of role of bacteria and extracellular polymeric substances in enhancing flocculating activity of algae (Grossart et al., 2006b; Kim et al., 2011; Tolhursf et al., 2002). However, the bacteria involved in flocculation and mechanism behind the process have been largely unclear. This study unveils the plausible role of algal-bacterial association in flocculation by experimenting with axenic, xenic and incomplete axenic (partially purified) cultures of Chlorella vulgaris under different ionic conditions. 2. Methods
2.3. Flocculation of axenic C. vulgaris cells mixed with microalgal cellfree xenic culture broth Two kinds of xenic culture broth was prepared after removing microalgal cells (bacterial cell broth) and all microbial cells (filtered broth) from the culture of xenic C. vulgaris. In the bacterial cell broth, only microalgal cells were removed by weak centrifugation at 3515g for 10 min from xenic C. vulgaris culture leaving behind the bacterial cells, as the name suggests. In the filtered broth, all microbial cells were removed from xenic C. vulgaris culture by 0.22 lm Millipore Express PLUS membrane filter at <40 psi (Fig. 1B). The pH of each broth was adjusted to 11 with 1 N NaOH. For the flocculation analysis, 6 106 cells ml1 of C. vulgaris cells were taken from the axenic culture and then mixed with the bacterial cell broth and filtered broth, respectively. Flocculating activity and pH of axenic C. vulgaris cells mixed with bacterial cell broth and filtered broth were measured immediately and after incubation for 24 h at room temperature. 2.4. DGGE analysis and sequencing
2.1. Axenic and xenic culture of C. vulgaris C. vulgaris (NCBI accession number JQ664295) used in this study was obtained from swine wastewater in Gonju, Korea, and was grown in BG11 medium (UTEX, 2009). C. vulgaris was inoculated into 300 ml of the BG11 medium in 1 L Erlenmeyer flask. The algal culture was stirred at 100 rpm, at 25 °C with a light intensity of 100 lmol m2 s1. Axenic C. vulgaris colony was obtained in consequent treatment of ultrasonication, fluorescence activated cell sorter (FACS), and micropicking from swine wastewater sample (D-H.C., R.R., J.L., B-H.K., H-M.O., and H-S.K., unpublished data). Long-term laboratory xenic culture of C. vulgaris was maintained by routine serial subculture over 3 months. Each C. vulgaris culture was cultivated for 14 days in BG11 medium. 2.2. Optimization of pH, flocculant concentration and flocculating activity After cultivation, pH of the cultures of axenic and xenic C. vulgaris was 8.1 and 8.9, respectively. The cultures were diluted with BG11 to equalize the microalgal cell concentrations to 6 106 cells ml1, and the pH was adjusted to 3, 5, 7, 9 and 11 with 1 N NaOH and 1 N HCl. 50 ml of C. vulgaris culture was mixed rapidly (300 rpm) for 30 s followed by slow mixing at 100 rpm for 1 min. Subsequently, cationic coagulant, CaCl2 (10 mM) was added followed by rapid mixing at 300 rpm for 30 s and slow mixing at 100 rpm for 1 min. Then FeCl3 (0.26 mM) was added to the culture, and mixed once again rapidly at 300 rpm for 30 s and slowly at 100 rpm for 1 min (Fig. 1A). The cultures were left for 2 min and 1 ml of aliquot was withdrawn and cell number was counted using C-chip hemocytometer (Digital Bio, Korea) at 200 magnification in an optical microscope (Nikon, Japan). Flocculating activity was calculated by the following equation (Kim et al., 2011; Oh et al., 2001):
A Flocculating activity ð%Þ ¼ 1 100 B
where A is the cell number after flocculation and B is the cell number before flocculation. The flocculating activity was also monitored at different concentrations of the flocculants used in this study, CaCl2 and FeCl3, respectively. At the end of flocculation experiment, cells were observed on a microscope to check for the differences in floc morphology (Microphot-FXA, Nikon, Japan). The zeta potential was measured in folded capillary cells with undiluted 1 ml samples of axenic and xenic culture (Zetasizer Nano ZS, Malvern, GK) (Henderson et al., 2008).
ð1Þ
Four different kinds of samples were used for extraction of genomic DNA: (1) Long-term laboratory xenic culture of C. vulgaris, (2) incomplete axenic culture, (3) supernatant of xenic C. vulgaris culture medium after centrifugation and (4) filtered xenic C. vulgaris culture medium. For denaturing gradient gel electrophoresis (DGGE) analysis, 16S rRNA gene sequence was partially amplified. PCR was performed with two sets of primers as follows; 341F with a GC-clamp (341F, 50 -CCT ACG GGA GGC AGC AG-30 ; GC-clamp 50 -CGC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GGG G-30 ) and 786R (50 -CTA CCA GGG TAT CTA ATC-30 ). DGGE was performed using the Dcode™ system (Bio-Rad Laboratories, USA). PCR products were applied directly onto 8% (w/v) polyacrylamide (37.5:1 acrylamide/bisacrylamide) gels with denaturing gradient range from 30% to 60% (100% denaturant solution contains 7 M urea and 40% v/v formamide). Electrophoresis was run in 1 TAE (40 mM Tris, 20 mM acetate, 1 mM EDTA pH 7.4) at 60 V at 60 °C for 18 h. Gels were stained with ethidium bromide (0.2 lg ml1, 1 TAE) for 15 min and destained in deionized water for 5 min and then visualized by using a KODAK Gel Logic 100 Imaging System. Each DGGE bands of interest was excised from the gels and cut bands were amplified as template for PCR. Forward and reverse strands sequences were assembled with SeqMan software (DNA STAR, Madison, WI) and homology searches of these assembled sequences were performed with the GenBank database using the Basic Local Alignment Search Tool (BLAST) in the NCBI (http://www.ncbi.nlm.nih.gov/). The sequences obtained in this study were deposited in the GenBank database under accession numbers from JX270632 to JX270636. 2.5. FACS treatment of xenic culture Incomplete axenic C. vulgaris was a partially purified xenic culture obtained by cell sorting using a BD FACSAria cell sorter (Becton Dickinson, USA). Flow-cytometry and cell sorting experiments
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
197
Fig. 1. (A). Flowchart of flocculation experiments performed in this study. 1 ml of aliquot was withdrawn near the water surface and cell number was counted by hemocytometer. (B) Different culture broths were used to determine the role of bacteria in flocculation.
were performed as mentioned in earlier studies (Doan et al., 2011; Sensena et al., 1993). Forward scatter width profiles were used to separate microalgal and bacterial cells. Microalgal cells were isolated through a 695/40 nm PerCP-Cy5.5 filter. The cells were sorted in a 15 ml centrifuge tubes containing sterile, half-strength BG11 medium. To establish clonal cultures, a single drop containing one cell was sorted into the tube. The tubes were opened just before the sorting event and closed immediately after delivering the drops. The sorting was conducted at least twice. The tubes were then cultured at 25 °C with continuous light intensity of 100 lmol/m1s1 of photons. The incomplete axenic cells were subjected to flocculation studies as mentioned earlier. 3. Results and discussion 3.1. Flocculating of axenic and xenic C. vulgaris cells and effect of pH on the flocculation Initially, flocculating activities of axenic and xenic cultures of C. vulgaris were compared at different flocculant concentrations (Fig. 2A and B) and in a pH range of 3–11 (Table 1). While no flocculating activity was observed in axenic C. vulgaris, long-term laboratory xenic culture of C. vulgaris showed high flocculating activities of more than 43% (±2.2) in the wide pH range of 3–11 with the highest activity of 94% (±4.9) at pH 11 (Table 1). These initial observations suggested that bacteria played a definite role in flocculation of microalgae. Moreover, results demonstrated that flocculation of xenic microalgal cells increased as pH and flocculant
concentration increased (Fig. 2A and B, Table 1). However, change in pH and flocculants did not flocculate axenic cultures. These result taken together suggested that flocculation of C. vulgaris was directly dependent on the associated bacterial communities. To study the role of bacterial communities further, a microscopic examination of axenic and xenic cultures after flocculation was undertaken (Fig. 3A and B). The results revealed that both axenic and xenic cultures formed flocs, however, size of flocs differed significantly. While the size of axenic flocs were about 20 lm or lesser, the size of xenic flocs were about 100 lm or more resulting in higher sedimentation and hence higher flocculating activity. These results pointed to the role of bacteria in formation of larger flocs, i.e. in floc growth. Hence to confirm these findings, studies on zeta potential of axenic and xenic cultures were performed under optimum conditions. Interestingly, the zeta potential measurements of axenic cultures were close to zero, which demonstrated the perfect stability of the flocs (Fig. 4A). On the contrary, the zeta potential values of xenic culture was about 15, which suggest inadequate stability of the flocs (Fig. 4B), however, the flocculating activity of axenic and xenic cultures were 2.1% and 94%, respectively at pH 11. A zeta potential value of zero suggests perfect floc formation, resultant from charge neutralization and definite aggregation of individual particles in a suspension (Henderson et al., 2008; Vallar et al., 1999). Both microscopic examination and zeta potential experiments clearly demonstrate that floc size influences the sedimentation characteristics of microalgal cell aggregates than the actual nature of the floc. Moreover, the role of bacteria in the formation of flocs is
198
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
Fig. 3. Light microscopic photographs of flocculated axenic (A) and xenic (B) cultures of C. vulgaris under optimum conditions.
3.2. Flocculation of axenic C. vulgaris cells mixed with microalgal cellfree xenic culture broth
Fig. 2. A three dimensional plot of flocculating activity of axenic (A) and xenic (B) cultures of C. vulgaris under differing CaCl2 and FeCl3 concentrations. Data were determined in duplicates.
also noteworthy. These results also point to two types of microalgal floc formation under gravity sedimentation, i.e. settleable flocs (in xenic cultures) and non-settleable flocs (in axenic cultures). Similar studies on role of algal-bacterial association in flocculation, especially in diatoms, suggest the role of light and production of transparent exopolymer particles as the glue for particle aggregation and floc formation (Gardes et al., 2011; Grossart et al., 2006a). In this study, increasing pH seems to confer an overall negative charge on both algae and associated bacteria, resulting in more effective adhesion during impact with flocculants. However, further studies were conducted to ascertain whether bacteria or its extracellular metabolites played the direct role in floc formation.
Further flocculation studies with two kinds of xenic culture broth, bacterial cell broth and filtered broth, were conducted to understand the source of flocculation. Compared with the flocculating activity of axenic culture, broth treated xenic culture showed higher flocculating activity of 83% with bacterial cell broth and 55% with filtered broth, respectively at the day 0. Even after 24 h incubation, the axenic C. vulgaris cells showed no flocculating activity, but flocculating activities of broth treated cultures increased mildly (Fig. 5), whereas the pH decreased. Interestingly, when axenic C. vulgaris cells were mixed with bacterial cell broth, flocculating activity similar to the original xenic culture was obtained. Although the flocculating activity of filtered broth was 30% lower than bacterial cell broth, this value is over 50%, and much higher than that of axenic culture. This conclusively proved that both bacterial cell and some bacterial extracellular metabolites play a major role in flocculation of C. vulgaris. Nevertheless, bacteria as a whole might play a more profound role in flocculation than bacterial extracellular metabolites alone. From the results of this study, there is direct correlation between settleable floc formation and presence of microalgae-associated bacteria and its extracellular metabolites. Recent studies have also highlighted the importance of algal-bacterial consortium in flocculation (Guo et al., 2011).
Table 1 Effect of pH on flocculating activity of axenic, incomplete axenic, and xenic C. vulgaris cultures.
Axenic Incomplete axenic Xenic
pH 3
pH 5
pH 7
pH 9
pH 11
0 ± 2.3 0 ± 1.3 43.0 ± 2.2
0 ± 0.1 2.3 ± 0.5 68.1 ± 6.0
0 ± 4.4 2.5 ± 0.9 78.3 ± 5.2
0 ± 1.8 2.8 ± 1.3 69.6 ± 2.2
2.1 ± 0.8 0 ± 2.0 94.1 ± 4.9
Axenic, incomplete axenic [partially purified xenic cultures by FACS (95% of bacteria was removed)], and xenic C. vulgaris cultures were cultivated for 14 days. Flocculating activity was measured with CaCl2 and FeCl3 as described in Section 2. Values are given as mean (%) ± standard deviation from three samples. Algal cell was adjusted to 6 106 cells ml1 in all samples, and bacterial cell number was 7 105 cells ml1 in xenic culture.
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
199
Fig. 4. Flocculating activity and zeta potential measurements of axenic (A) and xenic (B) cultures of C. vulgaris under different pH and flocculant concentrations. Fig. 6. Analysis of bacterial diversity of various xenic culture broths of C. vulgaris used in this study. (A) xenic culture broth; (B) bacterial cell broth; (C) filtered broth. See Section 2 for nomenclature.
(Doan et al., 2011; Sensena et al., 1993). As revealed in Figs. 6 and 7, xenic culture showed six different bacterial bands on DGGE gel. However, in the FACS treated cultures (incomplete axenic cultures), only two bands (band numbers 5 and 6) were visible (Fig. 7) implying that other bacterial bands present in xenic culture (band numbers 1–4) were removed by FACS treatment. 3.5. Flocculation studies with incomplete axenic C. vulgaris culture
Fig. 5. Flocculating activity of axenic C. vulgaris cells in the xenic culture broths. The pH of all treatments was adjusted to 11. The activities were measured immediately after mixing microalgal cells with the xenic broths (0 h) and after incubation for 24 h at room temperature. Data were determined in duplicates.
The direct role of bacteria in flocculating microalgae shows much promise, especially in the endeavor towards auto-flocculation using this association.
The incomplete axenic cultures obtained by FACS treatment were subjected to further flocculation studies. Interestingly, incomplete axenic strain showed very little flocculating activity (less than 3%) similar to that of complete axenic culture (Table 1). These results emphasize that the bacterial species representing DGGE bands 1–4 (Fig. 4) could be largely involved in the flocculation of C. vulgaris. 3.6. Identification of algae-associated bacteria
3.3. Analysis of bacterial diversity in bacterial cell broth and filtered broth of xenic C. vulgaris by DGGE Since bacteria was found to have a profound effect on microalgal flocculation, DGGE was performed to identify the bacterial community involved in treated xenic culture broths and original xenic culture (Fig. 6). Bands of the original xenic culture and bacterial cell broth displayed identical pattern and intensity with six distinct bands, but no bands were visible in filtered broth. 3.4. FACS treatment of xenic C. vulgaris and DGGE analysis In order to identify the communities which were responsible for flocculation, the xenic culture of C. vulgaris was treated using FACS
The 16S rRNA gene sequences of the DGGE bands and their NCBI nucleotide BLAST results were obtained. The first two hits with highest similarity in the BLAST analysis are presented in Table 2. Bacterial communities identified by DGGE analysis include Flavobacterium sp. (band number 1, accession number JX270632), Terrimonas sp. (band number 2, accession number JX270633), Sphingobacterium sp. (band numbers 3 and 4, accession number JX270634), Rhizobium sp. (band number 5, accession number JX270635) and Hyphomonas sp. (band number 6, accession number JX270636). FACS treatment of xenic culture resulting in elimination of bacteria to create an incomplete axenic culture demonstrates that Flavobacterium sp., Terrimonas sp., and Sphingobacterium sp. are
200
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
1978). Recent studies on PGPR revealed that this PGPR significantly enhances microalgal growth (Hernandez et al., 2009). Although various studies on biofilm formation by two different species of Hyphomonas are conclusive (Langille et al., 2000; Quintero and Weiner, 1995), our results suggest that Hyphomonas sp. found in DGGE band 6 has no perceivable impact on the flocculation of C. vulgaris. Aggregation of algae not only depends on the associated bacteria but also on quality of its secreted components (Grossart et al., 2006a). Induced aggregation due to EPS secreted by microorganisms such as algae, bacteria do settle down the aggregation of the particle (Stoll and Buffle, 1996). The results from this study imply that bacterial metabolites play a considerable role in flocculation. However, their flocculating activity is slightly lower (about 30%) when the associated bacteria are avoided (Fig. 5). This suggests a wider role of microalgae-associated bacteria in flocculation of C. vulgaris. Among the bacteria identified to be significant by FACS treatment and subsequent DGGE analysis, Flavobacterium sp. has been proved as a floc-forming bacteria. In activated sludge process, Flavobacterium sp. showed good flocculant growth in the presence of both calcium and magnesium ions. Flavobacterium sp. seem to promote aggregation of negatively charged surfaces of adjacent cells and cations (Tezuka, 1969). According to Schäfer et al. (2002), Bacteroidetes such as Flavobacteria and Sphingobacteria also associate with diatoms. Recent study also proposed Sphingobacterium sp. as a bioflocculant (Bao-Tian et al., 2006). To the best of our knowledge, Terrimonas sp. has not been reported earlier for its flocculating ability. Although the role of Terrimonas sp. individually would not be ascertained in this study, collectively with Flavobacterium sp. and Sphingobacterium sp. Terrimonas sp. may positively affect floc formation of microalgae. 4. Conclusions
Fig. 7. DGGE analysis of xenic and incomplete axenic C. vulgaris. (A) long-term laboratory xenic culture of C. vulgaris; (B) in complete axenic C. vulgaris after treatment with FACS; Data were determined in duplicates. The prominent bands have been numbered and identified by sequencing (see Table 2).
Table 2 Sequence identities of DGGE bands and those of closet relative in the database. Similarity (%)
Three microalgae-associated bacteria, i.e. Flavobacterium, Terrimonas, and Sphingobacterium, and their metabolites have a collective role in the harvest of C. vulgaris. These bacteria, in fact, play the defining role in bigger floc formation resulting in settleable flocs, which ultimately bolsters microalgal harvest. The role of pH and flocculants in enhancing flocculation can be also inferred from this study. However, axenic C. vulgaris is not capable of flocculation even in the presence of flocculants and higher pH because of insufficient floc size resulting in non-settleable flocs. Therefore, this study conclusively demonstrates the role of bacteria in flocculation of C. vulgaris.
Band no.
Accession no.
Best match in Blast analysis
1
HE997061 JN657227
Flavobacterium sp. Jyi-05 16S rRNA gene Flavobacterium sp. EC4 16S rRNA gene
97 96
2
JX511969 JF803808
Terrimonas ferruginea 16S rRNA gene Terrimonas sp. M-8 16S rRNA gene
99 99
Acknowledgements
3
AM411962 AM411963
Sphingobacterium sp. P-38 16S rRNA gene Sphingobacterium sp. P-17 16S rRNA gene
95 95
4
AM411962 AM411963
Sphingobacterium sp. P-38 16S rRNA gene Sphingobacterium sp. P-17 16S rRNA gene
97 97
5
EU781656 AF316615
Rhizobium sp. W3 16S rRNA gene Agrobacterium albertimagni 16S rRNA gene
100 99
This research was supported by grants from the Korea Ministry of Education, Science and Technology as a ‘‘Global Frontier Project’’ (www.biomass.re.kr) and grant from the KRIBB (Korea Research Institute of Bioscience and Biotechnology) Research Initiative Program (www.kribb.re.kr).
6
AM990830 AB362259
Hyphomonas sp. MOLA 55 16S rRNA gene Hyphomonas polymorpha 16S rRNA gene
94 94
vital organisms involved in the flocculation of microalgae. The other two bacteria, Rhizobium sp. and Hyphomonas sp. might not have significant role in flocculation (Table 1 and Fig. 4), however, their role as growth enhancing bacteria cannot be ruled out. Rhizobium sp., is a part of a community of bacteria which promote plant growth and are collectively termed as Plant Growth Promoting Rhizobacteria (PGPR) (Kloepper et al., 1980; Kloepper and Schroth,
References Bao-Tian, S., Ai-Hua, Z., Sheng-Kang, L., Xiu-Lin, W., 2006. Composition and characteristics of a bioflocculant produced by a Sphingobacterium sp. Periodical of Ocean University of China 5 (1672–5174). Benemann, J.R., Weissman, J.C., Koopman, B.L., Oswald, W.J., 1977. Energy production by microbial photosynthesis. Nature 268, 19–23. Brennan, L., Owende, P., 2010. Biofuels from microalgae—a review of technologies for production, processing, and extractions of biofuels and co-products. Renewable and Sustainable Energy Reviews 14 (2), 557–577. Divakaran, R., Sivasankara Pillai, V.N., 2002. Flocculation of algae using chitosan. Journal of Applied Phycology 14 (5), 419–422. Doan, T.T.Y., Sivaloganathan, B., Obbard, J.P., 2011. Screening of marine microalgae for biodiesel feedstock. Biomass and Bioenergy 35 (7), 2534–2544.
J. Lee et al. / Bioresource Technology 131 (2013) 195–201 Ebeling, J.M., Sibrell, P.L., Ogden, S.R., Summerfelt, S.T., 2003. Evaluation of chemical coagulation–flocculation aids for the removal of suspended solids and phosphorus from intensive recirculating aquaculture effluent discharge. Aquacultural Engineering 29 (1–2), 23–42. Gardes, A., Iversen, M.H., Grossart, H.-P., Passow, U., Ullrich, M.S., 2011. Diatomassociated bacteria are required for aggregation of Thalassiosira weissflogii. ISME Journal 5 (3), 436–445. Grossart, H.-P., Czub, G., Simon, M., 2006a. Algae–bacteria interactions and their effects on aggregation and organic matter flux in the sea. Environmental Microbiology 8 (6), 1074–1084. Grossart, H.P., Kiørboe, T., Tang, K.W., Allgaier, M., Yam, E.M., Ploug, H., 2006b. Interactions between marine snow and heterotrophic bacteria: aggregate formation and microbial dynamics. Aquatic Microbial Ecology 42 (1), 19–26. Gudin, C., Thepenier, C., 1986. Bioconversion of solar energy into organic chemicals by microalgae. Advances in Biotechnological Processes 6 (0736–2293), 73–110. Guo, F., Zhang, S.-H., Yu, X., Wei, B., 2011. Variations of both bacterial community and extracellular polymers: the inducements of increase of cell hydrophobicity from biofloc to aerobic granule sludge. Bioresource Technology 102 (11), 6421–6428. Henderson, R.K., Parsons, S.A., Jefferson, B., 2008. Successful removal of algae through the control of zeta potential. Separation Science and Technology 43 (7), 1653–1666. Hernandez, J.-P., de-Bashan, L.E., Rodriguez, D.J., Rodriguez, Y., Bashan, Y., 2009. Growth promotion of the freshwater microalga Chlorella vulgaris by the nitrogen-fixing, plant growth-promoting bacterium Bacillus pumilus from arid zone soils. European Journal of Soil Biology 45 (1), 88–93. Tolhursf, T.J., Gust, G., Paterson, D.M., 2002. The influence of an extracellular polymeric substance (EPS) on cohesive sediment stability. In: Johan, C.W., Cees, K. (Eds.), Proceedings in Marine Science, vol. 5. Elsevier. Kim, D.-G., La, H.-J., Ahn, C.-Y., Park, Y.-H., Oh, H.-M., 2011. Harvest of Scenedesmus sp. with bioflocculant and reuse of culture medium for subsequent high-density cultures. Bioresource Technology 102 (3), 3163–3168. Kloepper, J.W., Leong, J., Teintze, M., Schroth, M.N., 1980. Enhanced plant growth by siderophores produced by plant growth-promoting rhizobacteria. Nature 286 (5776), 885–886. Kloepper, J.W., Schroth, M.N., 1978. Plant growthpromoting rhizobacteria on radishes. In: Proceeding of the 4th international conference on plant pathogenic bacteria, INRA, Angers, France. Station de Pathologie Vegetale et Phytobacteriologie. pp. 879–882. Langille, S.E., Geesey, G.G., Weiner, R.M., 2000. Polysaccharide-specific probes inhibit adhesion of Hyphomonas rosenbergii strain VP-6 to hydrophilic surfaces. Journal of Industrial Microbiology & Biotechnology 25 (2), 81–85. Lee, S.J., Kim, S.-B., Kwon, G.-S., Yoon, B.-D., Oh, H.-M., 1998. Effects of harvesting method and growth stage on the flocculation of the green alga Botryococcus braunii. Letters in Applied Microbiology 27 (1), 14–18. Molina Grima, E., Belarbi, E.H., Acién Fernández, F.G., Robles Medina, A., Chisti, Y., 2003. Recovery of microalgal biomass and metabolites: process options and economics. Biotechnology Advances 20 (7–8), 491–515. Oh, H.-M., Lee, S.J., Park, M.-H., Kim, H.-S., Kim, H.-C., Yoon, J.-H., Kwon, G.-S., Yoon, B.-D., 2001. Harvesting of Chlorella vulgaris using a bioflocculant from Paenibacillus sp. AM49. Biotechnology Letters 23 (15), 1229–1234.
201
Oswald, W.J., Golueke, C.G., 1960. Biological transformation of solar energy. In: Wayne, W.U. (Ed.), Advances in Applied Microbiology, vol. 2. Academic Press, pp. 223–262. Pushparaj, B., Pelosi, E., Torzillo, G., Materassi, R., 1993. Microbial biomass recovery using a synthetic cationic polymer. Bioresource Technology 43 (1), 59–62. Quintero, E.J., Weiner, R.M., 1995. Evidence for the adhesive function of the exopolysaccharide of hyphomonas strain MHS-3 in its attachment to surfaces. Applied and Environmental Microbiology 61 (5), 1897–1903. Schäfer, H., Abbas, B., Witte, H., Muyzer, G., 2002. Genetic diversity of ‘satellite’ bacteria present in cultures of marine diatoms. FEMS Microbiology Ecology 42 (1), 25–35. Schlesinger, A., Eisenstadt, D., Bar-Gil, A., Carmely, H., Einbinder, S., Gressel, J., 2012. Inexpensive non-toxic flocculation of microalgae contradicts theories; overcoming a major hurdle to bulk algal production. Biotechnology Advances 30 (5), 1023–1030. Sensena, C.W., Heimanna, K., Melkoniana, M., 1993. The production of clonal and axenic cultures of microalgae using fluorescence-activated cell sorting. European Journal of Phycology 28 (2), 93–97. Spilling, K., Seppälä, J., Tamminen, T., 2011. Inducing autoflocculation in the diatom Phaeodactylum tricornutum through CO2 regulation. Journal of Applied Phycology 23 (6), 959–966. Stoll, S., Buffle, J., 1996. Computer simulation of bridging flocculation processes: the role of colloid to polymer concentration ratio on aggregation kinetics. Journal of Colloid and Interface Science 180 (2), 548–563. Sukenik, A., Shelef, G., 1984. Algal autoflocculation—verification and proposed mechanism. Biotechnology and Bioengineering 26 (2), 142–147. Tezuka, Y., 1969. Cation-dependent flocculation in a Flavobacterium species predominant in activated sludge. Applied Microbiology 17 (2), 222–226. Uduman, N., Qi, Y., Danquah, M.K., Forde, G.M., Hoadley, A., 2010. Dewatering of microalgal cultures: a major bottleneck to algae-based fuels. Journal of Renewable and Sustainable Energy 2 (1), 012701. UTEX, 2009. http://web.biosci.utexas.edu/utex/mediaDetail.aspx?mediaID=26. Vallar, S., Houivet, D., El Fallah, J., Kervadec, D., Haussonne, J.M., 1999. Oxide slurries stability and powders dispersion: optimization with zeta potential and rheological measurements. Journal of the European Ceramic Society 19 (6–7), 1017–1021. Vandamme, D., Foubert, I., Fraeye, I., Meesschaert, B., Muylaert, K., 2012. Flocculation of Chlorella vulgaris induced by high pH: role of magnesium and calcium and practical implications. Bioresource Technology 105, 114–119. Vandamme, D., Foubert, I., Meesschaert, B., Muylaert, K., 2010. Flocculation of microalgae using cationic starch. Journal of Applied Phycology 22 (4), 525–530. Vandamme, D., Pontes, S.C.V., Goiris, K., Foubert, I., Pinoy, L.J.J., Muylaert, K., 2011. Evaluation of electro-coagulation–flocculation for harvesting marine and freshwater microalgae. Biotechnology and Bioengineering 108 (10), 2320–2329. Wilde, E., Benemann, J., Weissman, J., Tillett, D., 1991. Cultivation of algae and nutrient removal in a waste heat utilization process. Journal of Applied Phycology 3 (2), 159–167. Wyatt, N.B., Gloe, L.M., Brady, P.V., Hewson, J.C., Grillet, A.M., Hankins, M.G., Pohl, P.I., 2012. Critical conditions for ferric chloride-induced flocculation of freshwater algae. Biotechnology and Bioengineering 109 (2), 493–501.
Contents lists available at SciVerse ScienceDirect
Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Microalgae-associated bacteria play a key role in the flocculation of Chlorella vulgaris Jimin Lee a,b, Dae-Hyun Cho a, Rishiram Ramanan a, Byung-Hyuk Kim a, Hee-Mock Oh a,b, Hee-Sik Kim a,b,⇑ a b
Environmental Biotechnology Research Center, Korea Research Institute of Bioscience and Biotechnology, 125 Gwahak-ro, Yuseong-gu, Daejeon 305-806, Republic of Korea University of Science and Technology, 217 Gajeong-ro, Yuseong-gu, Daejeon, Republic of Korea
h i g h l i g h t s " Flocculation of axenic and xenic cultures of Chlorella vulgaris implicates bacteria. " DGGE analysis indicates presence of five species of microalgae-associated bacteria. " FACS treatment of xenic culture implicates three bacterial species in flocculation. " Bacteria and its extracellular substances increase floc size. " This study proves that bacteria play a major role in flocculation of microalgae.
a r t i c l e
i n f o
Article history: Received 16 July 2012 Received in revised form 22 October 2012 Accepted 28 November 2012 Available online 8 December 2012 Keywords: Algae Flocculation Chlorella vulgaris Microalgae-associated bacteria
a b s t r a c t Flocculation is most preferred method for harvesting microalgae, however, the role of bacteria in microalgal flocculation process is still unknown. This study proves that bacteria play a profound role in flocculating by increasing the floc size resulting in sedimentation of microalgae. A flocculating activity of 94% was achieved with xenic Chlorella vulgaris culture as compared to 2% achieved with axenic culture. Denaturing gradient gel electrophoresis (DGGE) analysis of 16S rRNA gene of xenic C. vulgaris culture revealed the presence of Flavobacterium sp., Terrimonas sp., Sphingobacterium sp., Rhizobium sp. and Hyphomonas sp. as microalgae-associated bacteria. However when Flavobacterium, Terrimonas, Sphingobacterium were eliminated by fluorescence activated cell sorter (FACS), flocculating activity reduced to 3%. Further studies with cell free extracts also suggest that bacterial extracellular substances might also have a role in enhancing flocculation. We conclude that the collective presence of certain bacteria is the determining factor in flocculation of C. vulgaris. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Microalgae are now receiving extensive global attention as a potential source for biofuel production after being touted as an alternative fuel source some decades ago (Benemann et al., 1977; Oswald and Golueke, 1960). The production process of biodiesel using microalgal biomass includes cultivation, harvest, oil extraction, and conversion. Harvesting of microalgae is one of the most critical steps which involves separation of biomass from culture medium and contributes about 20–30% of the total biomass production cost (Gudin and Thepenier, 1986; Uduman et al., 2010). Harvesting is especially critical when the product of interest is a relatively low-value product like biodiesel (Vandamme et al., ⇑ Corresponding author at: Environmental Biotechnology Research Center, Korea Research Institute of Bioscience and Biotechnology, 125 Gwahak-ro, Yuseong-gu, Daejeon 305-806, Republic of Korea. Tel.: +82 42 860 4326; fax: +82 42 879 8103. E-mail address: [email protected] (H.-S. Kim). 0960-8524/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2012.11.130
2010). The major challenges in harvesting microalgae is because of small size of microalgal single cell (typically a few micrometer) and its low concentration in the culture medium (0.5–2 g L1) (Schlesinger et al., 2012; Vandamme et al., 2011). Most common harvesting methods include flocculation, gravity sedimentation, centrifugation, filtration and ultrafiltration, sometimes with an additional flocculation step or with a combination of flocculation–flotation. Centrifugation is one of the preferred methods for microalgal cell recovery because of effectiveness and rapidness but it requires energy thereby increasing operating costs (Molina Grima et al., 2003). Membrane replacement is the major cost involved in filtration method and this also depends on the concentration of microalgae (Wilde et al., 1991). One of the most potential methods to reduce cost and energy usage during harvesting is flocculation (Wyatt et al., 2012). Algal cells carry negative charge on the cell surface preventing aggregation because of repulsion; however the addition of cationic metal ions such as Ca2+ and Fe3+ can neutralize this charge, leading to the aggregation of cells.
196
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
Various flocculation methods result in higher particle sizes to enable gravity sedimentation, centrifugal recovery, and filtration (Molina Grima et al., 2003). The harvesting of microalgal cells by flocculation is more convenient process than contemporary methods such as centrifugation and filtration because it allows the treatment of large quantities of microalgal culture (Lee et al., 1998) and can be applied to a wide range of species (Pushparaj et al., 1993). Among the various flocculants, Aluminium sulphate (Alum) is most widely used for removal of algae, because of ease of use and application (Ebeling et al., 2003; Schlesinger et al., 2012). However, it cannot be applied over a wide pH range, moreover, floc size with alum when compared to ferric flocs is smaller resulting in ineffective sedimentation (Ebeling et al., 2003). Other cations such as calcium and magnesium also have a positive effect on flocculation in high pH (Vandamme et al., 2012). In addition, cationic polymers such as chitosan (Divakaran and Sivasankara Pillai, 2002), or alkalis such as NaOH have been used to achieve better flocculation (Brennan and Owende, 2010). On the contrary, various species of algae have been reported to auto-flocculate (Spilling et al., 2011; Sukenik and Shelef, 1984). There have been reports of role of bacteria and extracellular polymeric substances in enhancing flocculating activity of algae (Grossart et al., 2006b; Kim et al., 2011; Tolhursf et al., 2002). However, the bacteria involved in flocculation and mechanism behind the process have been largely unclear. This study unveils the plausible role of algal-bacterial association in flocculation by experimenting with axenic, xenic and incomplete axenic (partially purified) cultures of Chlorella vulgaris under different ionic conditions. 2. Methods
2.3. Flocculation of axenic C. vulgaris cells mixed with microalgal cellfree xenic culture broth Two kinds of xenic culture broth was prepared after removing microalgal cells (bacterial cell broth) and all microbial cells (filtered broth) from the culture of xenic C. vulgaris. In the bacterial cell broth, only microalgal cells were removed by weak centrifugation at 3515g for 10 min from xenic C. vulgaris culture leaving behind the bacterial cells, as the name suggests. In the filtered broth, all microbial cells were removed from xenic C. vulgaris culture by 0.22 lm Millipore Express PLUS membrane filter at <40 psi (Fig. 1B). The pH of each broth was adjusted to 11 with 1 N NaOH. For the flocculation analysis, 6 106 cells ml1 of C. vulgaris cells were taken from the axenic culture and then mixed with the bacterial cell broth and filtered broth, respectively. Flocculating activity and pH of axenic C. vulgaris cells mixed with bacterial cell broth and filtered broth were measured immediately and after incubation for 24 h at room temperature. 2.4. DGGE analysis and sequencing
2.1. Axenic and xenic culture of C. vulgaris C. vulgaris (NCBI accession number JQ664295) used in this study was obtained from swine wastewater in Gonju, Korea, and was grown in BG11 medium (UTEX, 2009). C. vulgaris was inoculated into 300 ml of the BG11 medium in 1 L Erlenmeyer flask. The algal culture was stirred at 100 rpm, at 25 °C with a light intensity of 100 lmol m2 s1. Axenic C. vulgaris colony was obtained in consequent treatment of ultrasonication, fluorescence activated cell sorter (FACS), and micropicking from swine wastewater sample (D-H.C., R.R., J.L., B-H.K., H-M.O., and H-S.K., unpublished data). Long-term laboratory xenic culture of C. vulgaris was maintained by routine serial subculture over 3 months. Each C. vulgaris culture was cultivated for 14 days in BG11 medium. 2.2. Optimization of pH, flocculant concentration and flocculating activity After cultivation, pH of the cultures of axenic and xenic C. vulgaris was 8.1 and 8.9, respectively. The cultures were diluted with BG11 to equalize the microalgal cell concentrations to 6 106 cells ml1, and the pH was adjusted to 3, 5, 7, 9 and 11 with 1 N NaOH and 1 N HCl. 50 ml of C. vulgaris culture was mixed rapidly (300 rpm) for 30 s followed by slow mixing at 100 rpm for 1 min. Subsequently, cationic coagulant, CaCl2 (10 mM) was added followed by rapid mixing at 300 rpm for 30 s and slow mixing at 100 rpm for 1 min. Then FeCl3 (0.26 mM) was added to the culture, and mixed once again rapidly at 300 rpm for 30 s and slowly at 100 rpm for 1 min (Fig. 1A). The cultures were left for 2 min and 1 ml of aliquot was withdrawn and cell number was counted using C-chip hemocytometer (Digital Bio, Korea) at 200 magnification in an optical microscope (Nikon, Japan). Flocculating activity was calculated by the following equation (Kim et al., 2011; Oh et al., 2001):
A Flocculating activity ð%Þ ¼ 1 100 B
where A is the cell number after flocculation and B is the cell number before flocculation. The flocculating activity was also monitored at different concentrations of the flocculants used in this study, CaCl2 and FeCl3, respectively. At the end of flocculation experiment, cells were observed on a microscope to check for the differences in floc morphology (Microphot-FXA, Nikon, Japan). The zeta potential was measured in folded capillary cells with undiluted 1 ml samples of axenic and xenic culture (Zetasizer Nano ZS, Malvern, GK) (Henderson et al., 2008).
ð1Þ
Four different kinds of samples were used for extraction of genomic DNA: (1) Long-term laboratory xenic culture of C. vulgaris, (2) incomplete axenic culture, (3) supernatant of xenic C. vulgaris culture medium after centrifugation and (4) filtered xenic C. vulgaris culture medium. For denaturing gradient gel electrophoresis (DGGE) analysis, 16S rRNA gene sequence was partially amplified. PCR was performed with two sets of primers as follows; 341F with a GC-clamp (341F, 50 -CCT ACG GGA GGC AGC AG-30 ; GC-clamp 50 -CGC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GGG G-30 ) and 786R (50 -CTA CCA GGG TAT CTA ATC-30 ). DGGE was performed using the Dcode™ system (Bio-Rad Laboratories, USA). PCR products were applied directly onto 8% (w/v) polyacrylamide (37.5:1 acrylamide/bisacrylamide) gels with denaturing gradient range from 30% to 60% (100% denaturant solution contains 7 M urea and 40% v/v formamide). Electrophoresis was run in 1 TAE (40 mM Tris, 20 mM acetate, 1 mM EDTA pH 7.4) at 60 V at 60 °C for 18 h. Gels were stained with ethidium bromide (0.2 lg ml1, 1 TAE) for 15 min and destained in deionized water for 5 min and then visualized by using a KODAK Gel Logic 100 Imaging System. Each DGGE bands of interest was excised from the gels and cut bands were amplified as template for PCR. Forward and reverse strands sequences were assembled with SeqMan software (DNA STAR, Madison, WI) and homology searches of these assembled sequences were performed with the GenBank database using the Basic Local Alignment Search Tool (BLAST) in the NCBI (http://www.ncbi.nlm.nih.gov/). The sequences obtained in this study were deposited in the GenBank database under accession numbers from JX270632 to JX270636. 2.5. FACS treatment of xenic culture Incomplete axenic C. vulgaris was a partially purified xenic culture obtained by cell sorting using a BD FACSAria cell sorter (Becton Dickinson, USA). Flow-cytometry and cell sorting experiments
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
197
Fig. 1. (A). Flowchart of flocculation experiments performed in this study. 1 ml of aliquot was withdrawn near the water surface and cell number was counted by hemocytometer. (B) Different culture broths were used to determine the role of bacteria in flocculation.
were performed as mentioned in earlier studies (Doan et al., 2011; Sensena et al., 1993). Forward scatter width profiles were used to separate microalgal and bacterial cells. Microalgal cells were isolated through a 695/40 nm PerCP-Cy5.5 filter. The cells were sorted in a 15 ml centrifuge tubes containing sterile, half-strength BG11 medium. To establish clonal cultures, a single drop containing one cell was sorted into the tube. The tubes were opened just before the sorting event and closed immediately after delivering the drops. The sorting was conducted at least twice. The tubes were then cultured at 25 °C with continuous light intensity of 100 lmol/m1s1 of photons. The incomplete axenic cells were subjected to flocculation studies as mentioned earlier. 3. Results and discussion 3.1. Flocculating of axenic and xenic C. vulgaris cells and effect of pH on the flocculation Initially, flocculating activities of axenic and xenic cultures of C. vulgaris were compared at different flocculant concentrations (Fig. 2A and B) and in a pH range of 3–11 (Table 1). While no flocculating activity was observed in axenic C. vulgaris, long-term laboratory xenic culture of C. vulgaris showed high flocculating activities of more than 43% (±2.2) in the wide pH range of 3–11 with the highest activity of 94% (±4.9) at pH 11 (Table 1). These initial observations suggested that bacteria played a definite role in flocculation of microalgae. Moreover, results demonstrated that flocculation of xenic microalgal cells increased as pH and flocculant
concentration increased (Fig. 2A and B, Table 1). However, change in pH and flocculants did not flocculate axenic cultures. These result taken together suggested that flocculation of C. vulgaris was directly dependent on the associated bacterial communities. To study the role of bacterial communities further, a microscopic examination of axenic and xenic cultures after flocculation was undertaken (Fig. 3A and B). The results revealed that both axenic and xenic cultures formed flocs, however, size of flocs differed significantly. While the size of axenic flocs were about 20 lm or lesser, the size of xenic flocs were about 100 lm or more resulting in higher sedimentation and hence higher flocculating activity. These results pointed to the role of bacteria in formation of larger flocs, i.e. in floc growth. Hence to confirm these findings, studies on zeta potential of axenic and xenic cultures were performed under optimum conditions. Interestingly, the zeta potential measurements of axenic cultures were close to zero, which demonstrated the perfect stability of the flocs (Fig. 4A). On the contrary, the zeta potential values of xenic culture was about 15, which suggest inadequate stability of the flocs (Fig. 4B), however, the flocculating activity of axenic and xenic cultures were 2.1% and 94%, respectively at pH 11. A zeta potential value of zero suggests perfect floc formation, resultant from charge neutralization and definite aggregation of individual particles in a suspension (Henderson et al., 2008; Vallar et al., 1999). Both microscopic examination and zeta potential experiments clearly demonstrate that floc size influences the sedimentation characteristics of microalgal cell aggregates than the actual nature of the floc. Moreover, the role of bacteria in the formation of flocs is
198
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
Fig. 3. Light microscopic photographs of flocculated axenic (A) and xenic (B) cultures of C. vulgaris under optimum conditions.
3.2. Flocculation of axenic C. vulgaris cells mixed with microalgal cellfree xenic culture broth
Fig. 2. A three dimensional plot of flocculating activity of axenic (A) and xenic (B) cultures of C. vulgaris under differing CaCl2 and FeCl3 concentrations. Data were determined in duplicates.
also noteworthy. These results also point to two types of microalgal floc formation under gravity sedimentation, i.e. settleable flocs (in xenic cultures) and non-settleable flocs (in axenic cultures). Similar studies on role of algal-bacterial association in flocculation, especially in diatoms, suggest the role of light and production of transparent exopolymer particles as the glue for particle aggregation and floc formation (Gardes et al., 2011; Grossart et al., 2006a). In this study, increasing pH seems to confer an overall negative charge on both algae and associated bacteria, resulting in more effective adhesion during impact with flocculants. However, further studies were conducted to ascertain whether bacteria or its extracellular metabolites played the direct role in floc formation.
Further flocculation studies with two kinds of xenic culture broth, bacterial cell broth and filtered broth, were conducted to understand the source of flocculation. Compared with the flocculating activity of axenic culture, broth treated xenic culture showed higher flocculating activity of 83% with bacterial cell broth and 55% with filtered broth, respectively at the day 0. Even after 24 h incubation, the axenic C. vulgaris cells showed no flocculating activity, but flocculating activities of broth treated cultures increased mildly (Fig. 5), whereas the pH decreased. Interestingly, when axenic C. vulgaris cells were mixed with bacterial cell broth, flocculating activity similar to the original xenic culture was obtained. Although the flocculating activity of filtered broth was 30% lower than bacterial cell broth, this value is over 50%, and much higher than that of axenic culture. This conclusively proved that both bacterial cell and some bacterial extracellular metabolites play a major role in flocculation of C. vulgaris. Nevertheless, bacteria as a whole might play a more profound role in flocculation than bacterial extracellular metabolites alone. From the results of this study, there is direct correlation between settleable floc formation and presence of microalgae-associated bacteria and its extracellular metabolites. Recent studies have also highlighted the importance of algal-bacterial consortium in flocculation (Guo et al., 2011).
Table 1 Effect of pH on flocculating activity of axenic, incomplete axenic, and xenic C. vulgaris cultures.
Axenic Incomplete axenic Xenic
pH 3
pH 5
pH 7
pH 9
pH 11
0 ± 2.3 0 ± 1.3 43.0 ± 2.2
0 ± 0.1 2.3 ± 0.5 68.1 ± 6.0
0 ± 4.4 2.5 ± 0.9 78.3 ± 5.2
0 ± 1.8 2.8 ± 1.3 69.6 ± 2.2
2.1 ± 0.8 0 ± 2.0 94.1 ± 4.9
Axenic, incomplete axenic [partially purified xenic cultures by FACS (95% of bacteria was removed)], and xenic C. vulgaris cultures were cultivated for 14 days. Flocculating activity was measured with CaCl2 and FeCl3 as described in Section 2. Values are given as mean (%) ± standard deviation from three samples. Algal cell was adjusted to 6 106 cells ml1 in all samples, and bacterial cell number was 7 105 cells ml1 in xenic culture.
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
199
Fig. 4. Flocculating activity and zeta potential measurements of axenic (A) and xenic (B) cultures of C. vulgaris under different pH and flocculant concentrations. Fig. 6. Analysis of bacterial diversity of various xenic culture broths of C. vulgaris used in this study. (A) xenic culture broth; (B) bacterial cell broth; (C) filtered broth. See Section 2 for nomenclature.
(Doan et al., 2011; Sensena et al., 1993). As revealed in Figs. 6 and 7, xenic culture showed six different bacterial bands on DGGE gel. However, in the FACS treated cultures (incomplete axenic cultures), only two bands (band numbers 5 and 6) were visible (Fig. 7) implying that other bacterial bands present in xenic culture (band numbers 1–4) were removed by FACS treatment. 3.5. Flocculation studies with incomplete axenic C. vulgaris culture
Fig. 5. Flocculating activity of axenic C. vulgaris cells in the xenic culture broths. The pH of all treatments was adjusted to 11. The activities were measured immediately after mixing microalgal cells with the xenic broths (0 h) and after incubation for 24 h at room temperature. Data were determined in duplicates.
The direct role of bacteria in flocculating microalgae shows much promise, especially in the endeavor towards auto-flocculation using this association.
The incomplete axenic cultures obtained by FACS treatment were subjected to further flocculation studies. Interestingly, incomplete axenic strain showed very little flocculating activity (less than 3%) similar to that of complete axenic culture (Table 1). These results emphasize that the bacterial species representing DGGE bands 1–4 (Fig. 4) could be largely involved in the flocculation of C. vulgaris. 3.6. Identification of algae-associated bacteria
3.3. Analysis of bacterial diversity in bacterial cell broth and filtered broth of xenic C. vulgaris by DGGE Since bacteria was found to have a profound effect on microalgal flocculation, DGGE was performed to identify the bacterial community involved in treated xenic culture broths and original xenic culture (Fig. 6). Bands of the original xenic culture and bacterial cell broth displayed identical pattern and intensity with six distinct bands, but no bands were visible in filtered broth. 3.4. FACS treatment of xenic C. vulgaris and DGGE analysis In order to identify the communities which were responsible for flocculation, the xenic culture of C. vulgaris was treated using FACS
The 16S rRNA gene sequences of the DGGE bands and their NCBI nucleotide BLAST results were obtained. The first two hits with highest similarity in the BLAST analysis are presented in Table 2. Bacterial communities identified by DGGE analysis include Flavobacterium sp. (band number 1, accession number JX270632), Terrimonas sp. (band number 2, accession number JX270633), Sphingobacterium sp. (band numbers 3 and 4, accession number JX270634), Rhizobium sp. (band number 5, accession number JX270635) and Hyphomonas sp. (band number 6, accession number JX270636). FACS treatment of xenic culture resulting in elimination of bacteria to create an incomplete axenic culture demonstrates that Flavobacterium sp., Terrimonas sp., and Sphingobacterium sp. are
200
J. Lee et al. / Bioresource Technology 131 (2013) 195–201
1978). Recent studies on PGPR revealed that this PGPR significantly enhances microalgal growth (Hernandez et al., 2009). Although various studies on biofilm formation by two different species of Hyphomonas are conclusive (Langille et al., 2000; Quintero and Weiner, 1995), our results suggest that Hyphomonas sp. found in DGGE band 6 has no perceivable impact on the flocculation of C. vulgaris. Aggregation of algae not only depends on the associated bacteria but also on quality of its secreted components (Grossart et al., 2006a). Induced aggregation due to EPS secreted by microorganisms such as algae, bacteria do settle down the aggregation of the particle (Stoll and Buffle, 1996). The results from this study imply that bacterial metabolites play a considerable role in flocculation. However, their flocculating activity is slightly lower (about 30%) when the associated bacteria are avoided (Fig. 5). This suggests a wider role of microalgae-associated bacteria in flocculation of C. vulgaris. Among the bacteria identified to be significant by FACS treatment and subsequent DGGE analysis, Flavobacterium sp. has been proved as a floc-forming bacteria. In activated sludge process, Flavobacterium sp. showed good flocculant growth in the presence of both calcium and magnesium ions. Flavobacterium sp. seem to promote aggregation of negatively charged surfaces of adjacent cells and cations (Tezuka, 1969). According to Schäfer et al. (2002), Bacteroidetes such as Flavobacteria and Sphingobacteria also associate with diatoms. Recent study also proposed Sphingobacterium sp. as a bioflocculant (Bao-Tian et al., 2006). To the best of our knowledge, Terrimonas sp. has not been reported earlier for its flocculating ability. Although the role of Terrimonas sp. individually would not be ascertained in this study, collectively with Flavobacterium sp. and Sphingobacterium sp. Terrimonas sp. may positively affect floc formation of microalgae. 4. Conclusions
Fig. 7. DGGE analysis of xenic and incomplete axenic C. vulgaris. (A) long-term laboratory xenic culture of C. vulgaris; (B) in complete axenic C. vulgaris after treatment with FACS; Data were determined in duplicates. The prominent bands have been numbered and identified by sequencing (see Table 2).
Table 2 Sequence identities of DGGE bands and those of closet relative in the database. Similarity (%)
Three microalgae-associated bacteria, i.e. Flavobacterium, Terrimonas, and Sphingobacterium, and their metabolites have a collective role in the harvest of C. vulgaris. These bacteria, in fact, play the defining role in bigger floc formation resulting in settleable flocs, which ultimately bolsters microalgal harvest. The role of pH and flocculants in enhancing flocculation can be also inferred from this study. However, axenic C. vulgaris is not capable of flocculation even in the presence of flocculants and higher pH because of insufficient floc size resulting in non-settleable flocs. Therefore, this study conclusively demonstrates the role of bacteria in flocculation of C. vulgaris.
Band no.
Accession no.
Best match in Blast analysis
1
HE997061 JN657227
Flavobacterium sp. Jyi-05 16S rRNA gene Flavobacterium sp. EC4 16S rRNA gene
97 96
2
JX511969 JF803808
Terrimonas ferruginea 16S rRNA gene Terrimonas sp. M-8 16S rRNA gene
99 99
Acknowledgements
3
AM411962 AM411963
Sphingobacterium sp. P-38 16S rRNA gene Sphingobacterium sp. P-17 16S rRNA gene
95 95
4
AM411962 AM411963
Sphingobacterium sp. P-38 16S rRNA gene Sphingobacterium sp. P-17 16S rRNA gene
97 97
5
EU781656 AF316615
Rhizobium sp. W3 16S rRNA gene Agrobacterium albertimagni 16S rRNA gene
100 99
This research was supported by grants from the Korea Ministry of Education, Science and Technology as a ‘‘Global Frontier Project’’ (www.biomass.re.kr) and grant from the KRIBB (Korea Research Institute of Bioscience and Biotechnology) Research Initiative Program (www.kribb.re.kr).
6
AM990830 AB362259
Hyphomonas sp. MOLA 55 16S rRNA gene Hyphomonas polymorpha 16S rRNA gene
94 94
vital organisms involved in the flocculation of microalgae. The other two bacteria, Rhizobium sp. and Hyphomonas sp. might not have significant role in flocculation (Table 1 and Fig. 4), however, their role as growth enhancing bacteria cannot be ruled out. Rhizobium sp., is a part of a community of bacteria which promote plant growth and are collectively termed as Plant Growth Promoting Rhizobacteria (PGPR) (Kloepper et al., 1980; Kloepper and Schroth,
References Bao-Tian, S., Ai-Hua, Z., Sheng-Kang, L., Xiu-Lin, W., 2006. Composition and characteristics of a bioflocculant produced by a Sphingobacterium sp. Periodical of Ocean University of China 5 (1672–5174). Benemann, J.R., Weissman, J.C., Koopman, B.L., Oswald, W.J., 1977. Energy production by microbial photosynthesis. Nature 268, 19–23. Brennan, L., Owende, P., 2010. Biofuels from microalgae—a review of technologies for production, processing, and extractions of biofuels and co-products. Renewable and Sustainable Energy Reviews 14 (2), 557–577. Divakaran, R., Sivasankara Pillai, V.N., 2002. Flocculation of algae using chitosan. Journal of Applied Phycology 14 (5), 419–422. Doan, T.T.Y., Sivaloganathan, B., Obbard, J.P., 2011. Screening of marine microalgae for biodiesel feedstock. Biomass and Bioenergy 35 (7), 2534–2544.
J. Lee et al. / Bioresource Technology 131 (2013) 195–201 Ebeling, J.M., Sibrell, P.L., Ogden, S.R., Summerfelt, S.T., 2003. Evaluation of chemical coagulation–flocculation aids for the removal of suspended solids and phosphorus from intensive recirculating aquaculture effluent discharge. Aquacultural Engineering 29 (1–2), 23–42. Gardes, A., Iversen, M.H., Grossart, H.-P., Passow, U., Ullrich, M.S., 2011. Diatomassociated bacteria are required for aggregation of Thalassiosira weissflogii. ISME Journal 5 (3), 436–445. Grossart, H.-P., Czub, G., Simon, M., 2006a. Algae–bacteria interactions and their effects on aggregation and organic matter flux in the sea. Environmental Microbiology 8 (6), 1074–1084. Grossart, H.P., Kiørboe, T., Tang, K.W., Allgaier, M., Yam, E.M., Ploug, H., 2006b. Interactions between marine snow and heterotrophic bacteria: aggregate formation and microbial dynamics. Aquatic Microbial Ecology 42 (1), 19–26. Gudin, C., Thepenier, C., 1986. Bioconversion of solar energy into organic chemicals by microalgae. Advances in Biotechnological Processes 6 (0736–2293), 73–110. Guo, F., Zhang, S.-H., Yu, X., Wei, B., 2011. Variations of both bacterial community and extracellular polymers: the inducements of increase of cell hydrophobicity from biofloc to aerobic granule sludge. Bioresource Technology 102 (11), 6421–6428. Henderson, R.K., Parsons, S.A., Jefferson, B., 2008. Successful removal of algae through the control of zeta potential. Separation Science and Technology 43 (7), 1653–1666. Hernandez, J.-P., de-Bashan, L.E., Rodriguez, D.J., Rodriguez, Y., Bashan, Y., 2009. Growth promotion of the freshwater microalga Chlorella vulgaris by the nitrogen-fixing, plant growth-promoting bacterium Bacillus pumilus from arid zone soils. European Journal of Soil Biology 45 (1), 88–93. Tolhursf, T.J., Gust, G., Paterson, D.M., 2002. The influence of an extracellular polymeric substance (EPS) on cohesive sediment stability. In: Johan, C.W., Cees, K. (Eds.), Proceedings in Marine Science, vol. 5. Elsevier. Kim, D.-G., La, H.-J., Ahn, C.-Y., Park, Y.-H., Oh, H.-M., 2011. Harvest of Scenedesmus sp. with bioflocculant and reuse of culture medium for subsequent high-density cultures. Bioresource Technology 102 (3), 3163–3168. Kloepper, J.W., Leong, J., Teintze, M., Schroth, M.N., 1980. Enhanced plant growth by siderophores produced by plant growth-promoting rhizobacteria. Nature 286 (5776), 885–886. Kloepper, J.W., Schroth, M.N., 1978. Plant growthpromoting rhizobacteria on radishes. In: Proceeding of the 4th international conference on plant pathogenic bacteria, INRA, Angers, France. Station de Pathologie Vegetale et Phytobacteriologie. pp. 879–882. Langille, S.E., Geesey, G.G., Weiner, R.M., 2000. Polysaccharide-specific probes inhibit adhesion of Hyphomonas rosenbergii strain VP-6 to hydrophilic surfaces. Journal of Industrial Microbiology & Biotechnology 25 (2), 81–85. Lee, S.J., Kim, S.-B., Kwon, G.-S., Yoon, B.-D., Oh, H.-M., 1998. Effects of harvesting method and growth stage on the flocculation of the green alga Botryococcus braunii. Letters in Applied Microbiology 27 (1), 14–18. Molina Grima, E., Belarbi, E.H., Acién Fernández, F.G., Robles Medina, A., Chisti, Y., 2003. Recovery of microalgal biomass and metabolites: process options and economics. Biotechnology Advances 20 (7–8), 491–515. Oh, H.-M., Lee, S.J., Park, M.-H., Kim, H.-S., Kim, H.-C., Yoon, J.-H., Kwon, G.-S., Yoon, B.-D., 2001. Harvesting of Chlorella vulgaris using a bioflocculant from Paenibacillus sp. AM49. Biotechnology Letters 23 (15), 1229–1234.
201
Oswald, W.J., Golueke, C.G., 1960. Biological transformation of solar energy. In: Wayne, W.U. (Ed.), Advances in Applied Microbiology, vol. 2. Academic Press, pp. 223–262. Pushparaj, B., Pelosi, E., Torzillo, G., Materassi, R., 1993. Microbial biomass recovery using a synthetic cationic polymer. Bioresource Technology 43 (1), 59–62. Quintero, E.J., Weiner, R.M., 1995. Evidence for the adhesive function of the exopolysaccharide of hyphomonas strain MHS-3 in its attachment to surfaces. Applied and Environmental Microbiology 61 (5), 1897–1903. Schäfer, H., Abbas, B., Witte, H., Muyzer, G., 2002. Genetic diversity of ‘satellite’ bacteria present in cultures of marine diatoms. FEMS Microbiology Ecology 42 (1), 25–35. Schlesinger, A., Eisenstadt, D., Bar-Gil, A., Carmely, H., Einbinder, S., Gressel, J., 2012. Inexpensive non-toxic flocculation of microalgae contradicts theories; overcoming a major hurdle to bulk algal production. Biotechnology Advances 30 (5), 1023–1030. Sensena, C.W., Heimanna, K., Melkoniana, M., 1993. The production of clonal and axenic cultures of microalgae using fluorescence-activated cell sorting. European Journal of Phycology 28 (2), 93–97. Spilling, K., Seppälä, J., Tamminen, T., 2011. Inducing autoflocculation in the diatom Phaeodactylum tricornutum through CO2 regulation. Journal of Applied Phycology 23 (6), 959–966. Stoll, S., Buffle, J., 1996. Computer simulation of bridging flocculation processes: the role of colloid to polymer concentration ratio on aggregation kinetics. Journal of Colloid and Interface Science 180 (2), 548–563. Sukenik, A., Shelef, G., 1984. Algal autoflocculation—verification and proposed mechanism. Biotechnology and Bioengineering 26 (2), 142–147. Tezuka, Y., 1969. Cation-dependent flocculation in a Flavobacterium species predominant in activated sludge. Applied Microbiology 17 (2), 222–226. Uduman, N., Qi, Y., Danquah, M.K., Forde, G.M., Hoadley, A., 2010. Dewatering of microalgal cultures: a major bottleneck to algae-based fuels. Journal of Renewable and Sustainable Energy 2 (1), 012701. UTEX, 2009. http://web.biosci.utexas.edu/utex/mediaDetail.aspx?mediaID=26. Vallar, S., Houivet, D., El Fallah, J., Kervadec, D., Haussonne, J.M., 1999. Oxide slurries stability and powders dispersion: optimization with zeta potential and rheological measurements. Journal of the European Ceramic Society 19 (6–7), 1017–1021. Vandamme, D., Foubert, I., Fraeye, I., Meesschaert, B., Muylaert, K., 2012. Flocculation of Chlorella vulgaris induced by high pH: role of magnesium and calcium and practical implications. Bioresource Technology 105, 114–119. Vandamme, D., Foubert, I., Meesschaert, B., Muylaert, K., 2010. Flocculation of microalgae using cationic starch. Journal of Applied Phycology 22 (4), 525–530. Vandamme, D., Pontes, S.C.V., Goiris, K., Foubert, I., Pinoy, L.J.J., Muylaert, K., 2011. Evaluation of electro-coagulation–flocculation for harvesting marine and freshwater microalgae. Biotechnology and Bioengineering 108 (10), 2320–2329. Wilde, E., Benemann, J., Weissman, J., Tillett, D., 1991. Cultivation of algae and nutrient removal in a waste heat utilization process. Journal of Applied Phycology 3 (2), 159–167. Wyatt, N.B., Gloe, L.M., Brady, P.V., Hewson, J.C., Grillet, A.M., Hankins, M.G., Pohl, P.I., 2012. Critical conditions for ferric chloride-induced flocculation of freshwater algae. Biotechnology and Bioengineering 109 (2), 493–501.