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Biochemical Engineering Journal 81 (2013) 170– 176

Contents lists available at ScienceDirect

Biochemical Engineering Journal

journa l h om epage: www.elsev ier .com/ locate /be j

egular Article

mproving microalgal oil collecting efficiency by pretreating theicroalgal cell wall with destructive bacteria

hun-Yen Chena, Ming-Der Baib, Jo-Shu Changa,c,d,∗

University Center for Bioscience and Biotechnology, National Cheng Kung University, Tainan 701, TaiwanGreen Energy and Environment Research Laboratories, Industrial Technology Research Institute, Hsinchu, TaiwanDepartment of Chemical Engineering, National Cheng Kung University, Tainan 701, TaiwanResearch Center for Energy Technology and Strategy, National Cheng Kung University, Tainan 701, Taiwan

r t i c l e i n f o

rticle history:eceived 27 July 2013eceived in revised form2 September 2013ccepted 17 October 2013vailable online 24 October 2013

eywords:icroalgae

a b s t r a c t

Converting microalgae oil into biodiesel is considered a promising route in the field of biofuel production.However, the cost of microalgae-based biodiesel is still too high to be economically feasible. The highcost of microalgae-based biodiesel is mainly a result of downstream processing, in particular from theextraction of oil out of the microalgal biomass. This study proposes a bacterial (enzymatic) destructionpretreatment of the microalgae cell wall to render the microalgal oil extraction more efficient. It has beenfound that the cell wall of microalgae can be modified if the microalgal biomass is co-cultured with anindigenous bacterial isolate Flammeovirga yaeyamensis in a salt concentration of 3% and a pH of 8.0. Fol-lowing this treatment, the activities of some hydrolytic enzymes (i.e., amylase, cellulases, and xylanase)

ell disruptionnzyme activitymylaseatch processing

have been detected in the co-culture of F. yaeyamensis and the oil-rich microalga (Chlorella vulgaris ESP-1). The SEM micrographs clearly show specific damage to the microalgae cell wall caused by the bacterialtreatment. We found that when the microalgae is pretreated with a concentrated co-culture supernatant(containing the hydrolytic enzymes), a nearly 100% increase in lipid extraction efficiency is obtained.The proposed bacterial disruption method seems to be an effective and environmentally friendly way ofimproving the efficiency of oil extraction and biofuel production from a microalgal biomass.

. Introduction

Research into the development of sustainable energy resourcesnd the reduction of carbon dioxide emissions is thriving due tooaring oil price and global climate change. Among the options forenewable energy, biofuels produced from biomass feedstock are ofost interest to the global energy structure [1,2]. Recently microal-

ae has received great attentions as a feedstock for biofuels due tots rapid growth compared to regular terrestrial plants and the effi-iency with which it captures carbon dioxide from the atmosphere3]. The growth of microalgae has few territorial restraints andhe resulting microalgal biomass contains valuable components,uch as proteins, sugars and lipids [4–6]. Moreover, lipid-deprivedesidues can also be used as a substrate in the production of

ther biofuels, such as hydrogen, bioethanol, and methane [7–10].herefore, microalgal biofuels seem to be very promising and arexpected to play a critical role in future energy development.

∗ Corresponding author at: Department of Chemical Engineering, National Chengung University, Tainan 710, Taiwan. Tel.: +886 6 2757575-62651;

ax: +886 6 2357146.E-mail address: changjs@mail.ncku.edu.tw (J.-S. Chang).

369-703X/$ – see front matter © 2013 Elsevier B.V. All rights reserved.ttp://dx.doi.org/10.1016/j.bej.2013.10.014

© 2013 Elsevier B.V. All rights reserved.

The key technology based requirements for the development ofmicroalgae-based biofuels include isolation of oleaginous microal-gae, establishment of effective microalgae cultivation strategies,and the effective execution of certain downstream processes, con-sisting of algal cell harvesting, cell disruption, lipid extraction,and the subsequent transesterification of microalgal lipids intobiodiesel [11–13]. Among these required steps, the cell disrup-tion treatment, which significantly influences the efficiency of oilextraction, is one of the most energy intensive steps in the overallbiodiesel synthesis process when using microalgae as feedstock.Therefore, finding an effective, environmentally friendly, and eco-nomically feasible method for the disruption of microalgal cellsis crucial for lowering the production cost of microalgae-basedbiodiesel.

Conventional cell disruption methods usually rely on physical orchemical treatments, which are either energy intensive or requirea large amount of hazardous chemicals, thereby adversely impact-ing the environment and significantly elevating the operating andmaintenance (O & M) costs. Some conventional cell disruption

methods may also denature the target products, such as pigments[14,15] and proteins [16,17]. In contrast, causing partial destructionof the microalgae cell wall through enzymatic hydrolysis has theadvantages of only requiring mild operation conditions, being high

gineering Journal 81 (2013) 170– 176 171

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C.-Y. Chen et al. / Biochemical En

elective in affect (e.g. no bad side-affects), and causing no damageo the extracted products [18]. It is also generally agreed that devel-ping enzymatic approaches helps facilitate the extraction of lipidsnd other commercially valuable byproducts from microalgae asell as reducing the quantity of solvents used, which is obviously

eneficial for the environment [19–22]. However, the structure andomposition of the algal cell walls may show some variation due tonvironmental and genetic differences [23,24], making enzymaticisruption of algal cell wall more difficult to consistently achieve.oreover, when using purified enzymes for the treatment, the cost

f the enzymatic disruption of the algal cell wall is often higher thanhemical or physical methods. Hence, a need has developed for aore economic method of biologically disrupting the cell wall, such

s using specialized bacteria.In this study, an indigenous bacterium (Flammeovirga yaeya-

ensis) [25] which secretes a specific digestive enzyme whichegrades the microalgae cell wall was co-cultured with an oleagi-ous green microalga Chlorella vulgaris ESP-31 [2]. The effect of theacterial disruption pretreatment on the cell wall was studied, andhe increased efficiency of algal lipid extraction was also evaluated.

. Materials and methods

.1. The microalgal strain and its cultivation conditions

The microalgal strain used in this work was isolated from ahrimp-culturing pond located in South Taiwan. The microalga wasdentified as C. vulgaris ESP-31 based upon its morphology as wells through 23S plastid rDNA sequence matching. The microalgatrain was grown on a Basal Medium [26] consisting of (g/l): KNO3,.25; KH2PO4, 1.25; MgSO4·7H2O, 1; CaCl2, 0.0835; H3BO3, 0.1142;eSO4·7H2O, 0.0498; ZnSO4·7H2O, 0.0882; MnCl2·4H2O, 0.0144;oO3, 0.0071; CuSO4·5H2O, 0.0157; Co(NO3)2·6H2O, 0.0049; EDTA

Na, 0.5. C. vulgaris ESP-31 was grown at 28 ◦C for 10 days with aontinuous supply of 2% CO2 at an aeration rate of 0.2 vvm underllumination with TL5 tungsten filament lamps at a light intensityf 60 �mol m−2 s−1. The total lipid content under those conditionsas estimated to be 21% per dry cell weight.

.2. The bacterial strain used for algal-cell-wall destruction

F. yaeyamensis was also isolated in southern Taiwan [25]. Theacterial strain was grown on an enriched YP agar plate (yeastxtract 2 g/l, peptone 20 g/l, sea salt 30 g/l and agar 15 g/l) or YProth (yeast extract 2 g/l, peptone 20 g/l, sea salt 30 g/l and agar

g/l) at a temperature of 30 ◦C.

.3. Co-culture of F. yaeyamensis and C. vulgaris ESP-31

The sterile modified f/2 medium [27] supplemented witheptone (2 g/l) and yeast extract (0.2 g/l) as additional nitrogenources was used to enhance the production of algal-cell-wallegrading enzymes from F. yaeyamensis [28]. For the co-culture,he microalgal biomass was first inoculated into the medium to anal concentration of 10 g/l. After that, F. yaeyamensis was inocu-

ated into the culture medium at an inoculums size of 0.1 g/l. Theo-culture was then kept at a temperature of 30 ◦C with an agitationate of 100 rpm for 10 days. During the course of co-culturing, theptical densities of the culture samples at 600 nm (OD600 nm) andt 688 nm (OD688 nm) were measured to estimate the concentration

f the bacterium and microalgae, respectively. Meanwhile, the pHalues, changes in sugar levels and enzyme activities (amylase, cel-ulase and xylanase) of the culture were also monitored during the0 days of cultivation.

Time (d )

Fig. 1. Effects of salt concentration on the growth of Flammeovirga yaeyamensis.

2.4. Measurement of light intensity

The light intensity on the reactor wall was measured using a LI-250 Light Meter with a LI-200SA pyranometer sensor (LI-COR, Inc.,Lincoln, Nebraska, USA). This light meter gives a measurement inunits of �mol m−2 s−1 for the light intensity.

2.5. Enzyme assay

The assay of the activity of cell-wall-degrading enzymes (i.e.,amylase, cellulases and xylanase) was based on the determinationof the amount of reducing sugars liberated from carboxymethyl-cellulose (CMC; purchased from Junsei), xylan (obtained fromSigma) and starch (obtained from Sigma), respectively. Amylaseactivity was analyzed after incubating 0.1 ml enzyme solution with0.5% starch in 100 mM potassium phosphate buffer (pH 7) at 37 ◦Cfor 30 min. Endo-glucanase activity was determined by incubating0.1 ml of supernatant with 0.9 ml of 1% carboxymethyl-cellulose(CMC) in McIlvaine’s buffer (pH 5), incubated at 50 ◦C for 30 min[29]. Xylanase activity was determined by incubating a 0.1 mlenzyme solution with 0.9 ml of 1% xylan in McIlvaine’s buffer (pH5), which was incubated at 55 ◦C for 10 min [29]. One unit of enzymeactivity (amylase, cellulases, and xylanase) was defined as theamount of enzyme required to produce 1.0 g/l of reducing sugarfrom the substrate per minute. Reducing sugar concentration wasdetermined by using the dinitrosalicylic acid (DNS) method [30].

2.6. Determination of the oil/lipids content

After achieving the desired level of cell growth, the microal-gae cells were harvested from culture broth using a centrifugation(9000 rpm for 10 min). The cells were washed twice with deionizedwater, lyophilized, and weighed. The microalgae sample (100 mg)was extract by 1:2 (v/v) chloroform and methanol at 200 rpm for30 min. After that, the same volume of chloroform was again addedto extract the lipids. Then, the same volume of water was added andthe same extraction procedures were repeated again. The resultingmixture was centrifuged at 1000 × g for 10 min and the chloroformphase was collected and subjected to evaporation under vacuum ina rotary evaporator to remove any remaining organic solvent. The

remnant from the evaporation was weighed as lipids.

The lipid composition was determined as fatty acid methylesters (FAMEs) through the direct transesterification method, asdeveloped by Lepage and Roy [31]. The C15 fatty acid was added

172 C.-Y. Chen et al. / Biochemical Engineering Journal 81 (2013) 170– 176

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Fig. 2. Effects of initial pH on the growth of Flammeovirga yaeyamensis.

s the internal standard to assist the quantification of the FAMEsontent. The sample was analyzed using gas chromatographyGC-2014, Shimadzu, Kyoto, Japan) equipped with a flame ioniza-ion detector (FID). Samples were then injected into a 30 m longapillary column (Type no. 260M142P, Thermo Fisher Scientific,

altham, MA, USA) with an internal diameter of 0.32 mm. Heliumas used as the carrier gas with a flow rate of 1.3 ml/min. The tem-eratures of the injector and detector were set at 250 and 280 ◦C,espectively. The oven temperature was initially set at 150 ◦C for.5 min, and then slowly increased, first to 180 ◦C at an increase ratef 10 ◦C/min, then to 220 ◦C with an increase rate of 1.5 ◦C/min, thenp to 260 ◦C with a 30 ◦C/min increase rate, and was finally held at60 ◦C for 5 min.

.7. SEM micrograph

The surface morphology of the microalgae cells before and afteracterial disruption was carefully examined using scanning elec-ron microscopy (SEM; Model S3000, Hitachi Co., Japan). Afterhe experiments, the algae cells were collected by centrifugation9000 rpm, 5 min). The collected cells were dehydrated in an ace-one solution for 30 min under a concentrations ranging from 50%o 95% (v/v) and then examined with a JEOL JSM-6700F scanninglectron microscope coupled with energy dispersive spectrometryEDS; Horiba, UK) [32].

. Results and discussion

.1. Effect of salt concentration and pH on the growth of F.aeyamensis

The effects of both the salt concentration and the pH value onhe growth of F. yaeyamensis used to disrupt the microalgal cell wallere investigated. Different salt concentrations (initial NaCl con-

entration = 1.0–6.0%) were used to cultivate F. yaeyamensis with aultivation temperature of 30 ◦C and an agitation rate of 100 rpm.

s indicated in Fig. 1, the optimal salt concentration for the growthf F. yaeyamensis was 3–4%, which corresponds well with its natu-al habitat, while a significant inhibition on F. yaeyamensis growthas observed when the salt content exceeded 5.0%. Consequently,

Fig. 3. Time course profile of (a) bacteria/microalgae ratios (OD600 nm/OD688 nm), (b)microalgae growth OD688 nm, (c) bacteria growth OD600 nm and (d) the reducing sugarconcentration in co-culture medium.

an initial salt concentration of 3.0% was determined to be ideal for

the growth of F. yaeyamensis.

The effect of the initial pH value on the growth of F. yaeyamensiswas also investigated. Fig. 2 shows the growth profile of F. yaeya-mensis at different starting pH levels. It shows that F. yaeyamensis

C.-Y. Chen et al. / Biochemical Engineering Journal 81 (2013) 170– 176 173

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rew fastest when the initial pH level was 6.0 and 7.0, while therowth rate decreased slightly at pH levels of 8.0 and 9.0. Nev-rtheless, the final cell concentration was similar for pH levelshroughout the whole range from 6.0–9.0. In contrast, when an ini-ial pH value of 10.0 was used, the growth of F. yaeyamensis wasuite poor with a significantly lower growth rate and final cell con-entration. Therefore, an initial pH of 6.0–7.0 is preferable for therowth of F. yaeyamensis.

.2. Co-culture of the bacterial and microalgal strains

In co-culture experiments, F. yaeyamensis was placed in an arti-cial sea water medium (modified f/2 medium) [27] to mimic theatural environment for bacterial growth. A small amount (10.0 g/l)f microalgae biomass was also added into the medium which hadlready been inoculated with F. yaeyamensis. Fig. 3a–c shows therowth index (OD600 nm/OD688 nm) of F. yaeyamensis. Absorbancet 600 nm (OD600 nm) indicates that the cell concentration in theedium and OD688 nm values represent unique absorbing pattern

f chlorophyll. Hence, a higher ratio of OD600 nm/OD688 nm leadso a higher rate of bacterial cell growth with a lower level ofon-damaged microalgae in the medium. The OD600 nm/OD688 nmatios were calculated from the slopes determined from measuringhe OD600 nm and OD688 nm values, respectively, at 1/40, 1/20 and/10 diluents for each day. As shown in Fig. 3a, the increase in theD600 nm/OD688 nm ratio over the time of cultivation clearly shows

he interrelated pattern of bacterial growth and the destruction of

he microalgal cells.

Further evidence showing the destruction and/or degradationf the microalgal cell wall was found by determining the reducingugar concentration released into the medium after the disruption

10 days with the co-culture of F. yaeyamensis and Chlorella vulgaris ESP-31.

of the microalgal cell wall. It is likely that the polysaccharides (in theform of starch and cellulose) were released into the extracellularcompartment during the breakdown of the cell wall. In addition,hydrolytic degradation of the polysaccharides with the enzymes(e.g., cellulases, amylases, etc.) produced by the cell-wall-degradingbacterium (i.e., F. yaeyamensis) may also produce soluble mono- oroligo-saccharides, which will be present in the supernatant of theco-culture medium, and can be measured as reducing sugars. Fig. 3dshows the amount of reducing sugars (RS) in the medium duringthe period of co-culture. The RS concentration was 0.2 g/l beforebacterial inoculation. From 1st to 10th day, the RS concentrationwas at a low level indicates the consumption of the released reduc-ing sugar (as the carbon source) during the bacterial growth. SEMmicrographs of microalgae were also captured at 10,000× mag-nification after co-culture with F. yaeyamensis (Fig. 4). The roughsurface of the microalgae indicates the decomposition of the algalcell wall after the bacterial treatment (Fig. 4). One of the major rea-sons for the damage to the microalgal cell wall is likely to be theenzymatic degradation, which has also been mentioned by otherresearchers [24]. The hypothesis of enzymatic destruction of themicroalgal cell wall was further confirmed by determining whetherthe enzymatic activities were indeed present in the co-culture. Thedetails of the enzyme assay are presented in the following section.

3.3. Activities of the hydrolytic enzymes in thebacterium/microalgae co-culture

Bacteria which can degrade the algal-cell-wall may secretedigesting enzymes to decompose the cell wall for its bacterialgrowth [28]. A series of enzymes are also produced and excretedfrom the cell-wall degrading bacteria when it senses it is near

174 C.-Y. Chen et al. / Biochemical Engineering Journal 81 (2013) 170– 176

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ig. 5. The profile of hydrolytic enzyme activities during the co-culture of F. yaeyylanase activity.

utrients, such as peptides, oligosaccharides and polysaccharides.he outer-layer composition and structure of microalgae may varyue to difference in microalgae species and environmental condi-ions [19,23,33]. In this study, the composition of carbohydratesn the microalga was preliminarily analyzed with high perfor-

ance liquid chromatography after acidic hydrolysis of microalgaliomass. It was found that the major sugar components of theicroalgae-based carbohydrates are glucose (mainly from cellu-

ose and starch), xylose, galactose, arabinose and rhamnose (dataot shown). Based on this analysis, the activities of the corre-ponding hydrolytic enzymes (such as amylase, endo-glucanasend xylanase) could be determined during the course of the co-ulture of F. yaeyamensis and C. vulgaris (Fig. 5a–c). One unit of each

nzyme activity was defined as the amount of enzyme which liber-ted 1 �mol reducing sugar per minute per milliliter. The amylasectivity was observed during the co-culture (Fig. 5a). Within threeays the amylase activity rapidly increased to nearly 2.5 U/ml and

is and C. vulgaris ESP-31. (a) Amylase activity, (b) endo-glucanase activity and (c)

eventually leveled off after day 6. The presence of amylase activitywas also found in other marine bacteria [34]. Activities of endo-glucanase (Fig. 5b) and xylanase (Fig. 5c) were also detected butthe activity was minor compared to that of amylase, especiallyfor endo-glucanase. The trends in the enzyme activity for amylaseand endo-glucanase were similar, as they both increased sharply inthe first 2–4 days and then leveled off or slightly decreased there-after. The lower enzyme activity for cellulose and xylanase might beattributed to less substrate stimulation [28]. In contrast, the amy-lase might be better expressed due to a greater availability of starchas the substrate in the medium.

3.4. Enzymatic destruction of microalgal cell wall

Cellulases are used to digest cellulose in straw and sugar-cane bagasse for biofuel production [35]. Since the inner cellwall of microalgae (e.g., C. vulgaris) is also mainly composed of

C.-Y. Chen et al. / Biochemical Engineering Journal 81 (2013) 170– 176 175

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Table 1The main fatty acid profile of the microalgae lipid.

Compound Percentage (%) per dry weight

Palmitic acid (C16:0) 6.64Palmitoleic acid (C16:1) 1.89Stearic acid (C18:0) 0.4Oleic acid (C18:1) 3.83Linoleic acid (C18:2) 7.96

10 g/l) using FY enzyme (containing 4 U/ml amylase, 5 mU/ml cellulase and 15 mUylanase), commercial cellulases (5 U/ml) and commercial amylase (5 U/ml). Con-rol: treatment of microalgal biomass with enzyme-free dionized water.

ignin-free cellulose, immobilized cellulases are utilized to degradehe microalgal cell wall for reducing sugar production and lipidxtraction from microalgae [36]. In this study, F. yaeyamensis wasound to secrete hydrolytic enzymes (such as amylases and cellu-ases) to degrade the microalgal cell wall. However, as shown inig. 5, the activities of endo-glucanase and xylanase in the bac-erium/microalga co-culture are much lower than that of amylase.n order to clarify which hydrolytic enzymes play the key role inhe degradation of the microalgal cell wall, a concentrated enzymeolution from the supernatant of 10-day co-culture of F. yaeya-ensis and C. vulgaris (denoted as FY enzyme containing 4 U/ml

mylase, 5 mU/ml cellulase and 15 mU xylanase), a solution ofommercial amylase (5 U/ml) (sigma # 10070-50G), and a solu-ion of commercial cellulase (5 U/ml) (sigma #C1184-5KU) weretilized for digesting microalgal cell wall for the comparison ofheir cell-wall degradation performance, which was justified byheir varying abilities in producing reducing sugar and varying effi-iency at promoting lipid extraction. Fig. 6 shows the reducingugar (RS) production for each period of treatment when using dif-erent types of enzymes. The maximum RS production was about.2 g/l, achieved when the sample was treated with FY enzyme orommercial cellulase alone, and was 0.15 g/l with commercial amy-ase alone. The RS production rate was markedly higher when usingommercial cellulase and FY enzymes rather than just using com-ercial amylase alone. This suggests that the enzymatic breakdown

f the cell wall through the hydrolytic degradation of cellulose onhe cell wall might be the rate-limiting step in the overall cell dis-uption process.

Fig. 7 shows algal oil/lipid extraction efficiency of the microalgaliomass undergoing different enzymatic treatments over differentime intervals (i.e., 0 h, 2 h, 4 h, 6 h, 1 day, and 3 days). The effi-iency of microalgal oil/lipid extraction in 3-day treatment withY enzyme is nearly 21.5%. This efficiency is nearly double that ofhe control (without enzymatic treatment) and was also 57–69%igher than that obtained from using commercial amylase and cel-

ulase alone (Fig. 7). Therefore, the treatment of the microalgalell wall with an appropriate combination of several hydrolyticnzymes (rather than with a single hydrolytic enzyme) seems toe required to generate the type of efficient cell disruption that is

Arachidic acid (C20:0) 1.14

able to enhance the efficiency of oil/lipid extraction. The resultsalso suggest that the cell wall destruction by the mixed enzymeswas a time consuming process (3 days in this case). A higherenzyme loading might be required to shorten the treatment time.A recent study also shows that immobilized commercial cellulasesare effective in digesting the microalgal cell wall and thus promot-ing reducing sugar production and oil extraction [36]. In addition,analysis of FAME composition of the extract microalgal lipid showsthat the main fatty acids components were palmitic acid (C16:0),palmitoleic acid (C16:1), stearic acid (C18:0), oleic acid (C18:1),linoleic acid (C18:2) and �-linolenic acid (C18:3) (Table 1). Thisstudy shows that a cocktail enzyme solution containing neededhydrolytic enzymes for microalgal cell wall degradation can be auseful tool for the cell wall treatment and achieves a better perfor-mance in regards to oil/lipid extraction as well as reducing sugarsproduction from microalgae.

4. Conclusion

This study shows that an isolated marine bacterium F. yaeya-mensis is able to effectively disrupt the cell wall of C. vulgaris via aco-culture of the bacterium in conjunction with microalga. One ofthe mechanisms leading to cell disruption is enzymatic hydrolysisof the cellulose components in the microalgal cell wall togetherwith the simultaneous hydrolysis of the starch content, result-ing in the liberation of reducing sugars. Treating the microalgalbiomass with concentrated enzymes derived from F. yaeyamen-sis (FY enzyme) seems to be significantly able in improving theoil/lipid extraction efficiency. This demonstrates the potential ofusing the present bacterial system for degrading the microalgae

cell wall to achieve a high-efficiency oil/lipid production from thetarget microalgae.

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cknowledgements

The authors gratefully acknowledge financial supports from thenergy and Environment Research Laboratories, Industrial Tech-ology Research Institute under grant numbers I550000534 andational Science Council of Taiwan (grant nos. NSC 101-3113-P-06-015, NSC 100-2221-E-006-126 and NSC 100-3113-E-006-016.he support from NCKU’s top university project (with 5-year-0-billion funding from Taiwan’s Ministry of Education) is alsoppreciated.

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