Bioresource Technology 102 (2011) 8191–8198

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Bioresource Technology

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Enhanced production of bioethanol and ultrastructural characteristics of reusedSaccharomyces cerevisiae immobilized calcium alginate beads

Kwang Ho Lee a,c, In Seong Choi a, Young-Gyu Kim b, Duck-Joo Yang c, Hyeun-Jong Bae b,d,⇑a Department of Wood Science and Landscape Architecture (BK21 Program), Chonnam National University, Gwangju 500-757, Republic of Koreab Department of Forest Products and Technology, Chonnam National University, Gwangju 500-757, Republic of Koreac Department of Chemistry and The AG MacDiarmid Nanotech Institute, The University of Texas at Dallas, TX 75080, USAd Department of Bioenergy Science and Technology, Chonnam National University, Gwangju 500-757, Republic of Korea

a r t i c l e i n f o a b s t r a c t

Article history:Received 29 March 2011Received in revised form 4 June 2011Accepted 16 June 2011Available online 23 June 2011

Keywords:Ethanol fermentationCalcium alginate beadsReused beadsUltrastructural characteristics, Egg-boxmodel

0960-8524/$ - see front matter � 2011 Elsevier Ltd. Adoi:10.1016/j.biortech.2011.06.063

⇑ Corresponding author. Address: Department of ForChonnam National University, Gwangju 500-757, Re530 2097; fax: +82 62 530 0029.

E-mail address: baehj@chonnam.ac.kr (H.-J. Bae).

Yeast immobilized on alginate beads produced a higher ethanol yield more rapidly than did free yeastcells under the same batch-fermentation conditions. The optimal fermentation conditions were 30 �C,pH 5.0, and 10% initial glucose concentration with 2% sodium alginate beads. The fermentation time usingreused alginate beads was 10–14 h, whereas fresh beads took 24 h, and free cells took 36 h. All bead sam-ples resulted in nearly a 100% ethanol yield, whereas the free cells resulted in an 88% yield. Transmissionelectron microscopy (TEM) showed that the shortened time and higher yield with the reused beads wasdue to a higher yeast population per bead as well as a higher porosity. The ultrastructure of calcium algi-nate beads and the alginate matrix structure known as the ‘‘egg-box’’ model were observed using TEM.

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1. Introduction

Interest in ethanol production from biomass feed stocks as analternative energy source from petroleum has grown as the priceof petroleum has generally increased over the last few years. Cur-rent research on bioethanol is driven by the need to reduce the costof ethanol production and has focused on improving feedstock pre-treatment methods, enzymes, and fermentation.

Ethanol is produced by four main types of industrial operations,including batch, fed-batch, continuous, and semi-continuous oper-ation (Çaylak, 1998). However, most ethanol produced today is pre-pared by batch operations. Because batch fermentation has theadvantage of low investment costs, simple controls, and operationsthat do not require specialized labor, complete sterilization andmanagement of feedstock are easier than in other processes(Çaylak, 1998). Another advantage of batch fermentation usingimmobilized yeast is the ability to separate immobilized yeast fromthe ethanol product, allowing the immobilized yeast to be reusedfor further fermentation (Bayramoglu and Arica, 2009). In additionto the smaller reactor size required, fermentation with immobilizedcells confers higher ethanol tolerance and cell concentrations,

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shorter fermentation time, increased periods of aging time andlong-term preservation, greater cell activity and stability, enhancedfermentation productivity, greater feasibility of continuousprocessing, lower costs of recovery and recycling, and ease of sepa-ration of the immobilized medium from the reaction (Kourkoutaset al., 2004; Tata et al., 1999).

Kourkoutas et al. (2004) discussed the properties of carriers andimmobilized cells. Immobilization techniques can be divided intofour main categories based on the physical mechanism employed:(1) attachment or adsorption on solid carrier surfaces; (2) entrap-ment within a porous matrix; (3) self aggregation by flocculation(natural) or with cross-linking agents (artificially induced); and (4)cell containment behind barriers. Various immobilization sub-strates have been used, including alumina beads and membranes(Hamdy et al., 1990), porous glass beads (Navarro and Durand,1977), diatomaceous earth (Navarro and Durand, 1977; Virkajärviand Pohjala, 2000), DEAE-cellulose (Virkajärvi and Pohjala, 2000),Ca-alginate beads (Kierstan and Bucke, 1997), j-carrageenan(Pilkington et al., 1999), wood chips (Vidgren et al., 2003), siliconcarbide (Tata et al., 1999), and spent grains (Brányik et al., 2001).

Extracting alginates from brown seaweed has been studied overseveral decades to develop economical processing, obtain highyields, determine the best conditions to handle the product, andestablish a controlled molecular weight for different applications(Gomez et al., 2009). Sodium alginate is a water-soluble polymerthat produces highly viscous solutions, a characteristic that

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contributes to stabilizing the suspension of microorganisms in thealginate matrix. Sodium alginate solutions form gels in the pres-ence of cations such as Ca2+ (Draget et al., 1997). Immobilizing cellsin alginate is simple, cheap, and non-toxic. Therefore, alginate isfrequently used for immobilization. Moreover, calcium alginate isthe most widely used gel matrix in laboratory, pilot-plant, andindustrial-scale fermentation projects (Ciesarová et al., 1998).

Ultrastructural characteristics of some immobilized matrixesand yeast cells such as Saccharomyces cerevisiae are immobilizedin hollow-fiber membrane bioreactors by Inloes et al. (1983) andin transmission electron microscopy (TEM) images of yeast byWright (2000). However, no studies have reported the ultrastruc-tural relationship between yeast cells and calcium alginate matri-ces within a calcium alginate bead. Calcium alginate beads treatedwith 100% ethanol shrink from 3.8 to 1.1 mm. Therefore, conven-tional pretreatment methods (chemical fixation and dehydrationin ethanol gradually increased to 100%, incubating cells with gentleagitation on a rotating drum mixer with several changes of resin:solvent mixtures, in which the concentration of resin gradually in-creases to 100%) cannot be applied for TEM observations of calciumalginate beads.

In this study, we confirmed that batch-type fermentation usingyeast immobilized in 2% calcium alginate beads with a 10% glucoseinitial concentration produced a higher ethanol yield than didbatch fermentation with mobile yeast. We identified the optimalfermentation conditions of temperature, pH and initial glucoseconcentration and the optimum sodium alginate concentration ofbeads. We also investigated the effect of reused alginate beadsvs. fresh beads on fermentation time and ethanol yield. This studyis the first observation of the ultrastructure of calcium alginatebeads and the alginate matrix structure known as the ‘‘egg-box’’model by TEM.

2. Methods

2.1. Preparation of culture medium

Saccharomyces cerevisiae KCTC 7906 (Biological Resource Cen-ter, Korea Research Institute of Bioscience and Biotechnology,Daejeon, Korea) cells were grown in media containing 5 g glucose,1 g peptone, 1 g yeast extract, 0.1 g MgSO4, and 0.1 g K2HPO4 in100 ml deionized water. The media was sterilized at 121 �C for20 min. At 30 �C, 0.6 g of yeast powder was added under sterileconditions. A seed culture of S. cerevisiae was grown in a shakingincubator at 30 �C and 300 rpm for 24 h. The yeast cell seed culturewas transferred to prepared fermentation-culture medium or usedfor making calcium alginate beads. Fermentation-culture mediumwas prepared using 100 g glucose for 10% (150 g glucose for15%), 5 g peptone, 5 g yeast extract, 1 g MgSO4, 1 g K2PO4 in 1 literdistilled water.

2.2. Preparation of calcium alginate beads

A 24-h culture was harvested at the exponential growth phaseand mixed with a sodium alginate solution. To prepare the calciumalginate beads, 100 ml of yeast seed culture was added to a sodiumalginate solution prepared by dissolving 8 g of powder for 2% so-dium alginate (Spectrum Chemical, Gardena, CA, USA) (10 g for2.5% sodium alginate) in 300 ml of deionized water (Najafpouret al., 2004). The mixture of sodium alginate and seed culturewas dropped through syringes into a 0.1 M CaCl2 solution. Thebeads were stored after washing with deionized water to removeresidual CaCl2.

The 3.8-mm beads were uniformly packed and stored in deion-ized water at 4 �C for 3 days before fermentation. Beads were

collected by filtration, added to the fermenter (Bioflo110 Fermen-ter/Bioreactor, New Brunswick Scientific, Edison, NJ, USA) with1 liter of fermentation medium at 300 rpm and varying tempera-tures, pHs, sodium alginate concentrations, and initial glucose con-centrations for each experiment. Samples were collected every 2 or3 h to determine the amount of residual glucose and the ethanolconcentration.

2.3. Glucose and ethanol concentration

Glucose concentration was measured using the dinitrosalicylicacid method (Miller, 1959). Ethanol concentrations were measuredby gas chromatography (GC-8A, Shimadzu, Tokyo, Japan) equippedwith a 20% carbowax column. Isopropanol was used as an internalstandard to measure ethanol content from each fermented sample.The column temperatures at the injector and detector were 130 �Cand 110 �C, respectively.

2.4. Ultrastructural characterization

Samples were taken from fresh beads and beads after 36 h fer-mentation in the batch fermenter for electron microscopy. Alginatebeads were fixed with a mixture of freshly prepared 2% glutaralde-hyde (v/v) and 2% paraformaldehyde (v/v) dissolved in 0.05 M cac-odylate buffer (pH 7.2) at room temperature for 4 h. After washingin the same buffer, samples were post-fixed with 1% osmium tetra-oxide in 0.05 M cacodylate buffer at room temperature for 1 h.After washing in the same buffer and dehydration with 30%, 50%,and 70% ethanol, the alginate beads were air dried at room temper-ature and examined under a scanning electron microscope (SEM;LEO 1530 VP) operated at 3 KV. Other samples for TEM observationwere fixed with the same fixative used for pretreating the SEMsamples and dehydrated with 30%, 50%, and 70% ethanol. Afterdehydration with 70% ethanol, the samples for resin infiltrationwere reacted with a mixture of 70% ethanol and London ResinWhite (LR White; London Resin, London, UK). After substitutionwith an ascending LR White series, the samples were embeddedin LR White. Ultrathin sections (80 nm) were prepared with anultramicrotome equipped with a diamond knife, mounted on un-coated nickel grids (300 mesh), and stained with 4% uranyl acetateand 1% lead citrate. The sections were examined with a JEM-1400(JEOL Ltd., Tokyo Japan) TEM at an acceleration voltage of 80 KV.

3. Results and discussion

3.1. Effect of sodium alginate concentration on glucose consumption

To determine the effect of sodium alginate concentration on fer-mentation yield, the residual glucose concentration after fermenta-tion with 2% and 2.5% calcium alginate beads or free cells (noimmobilized yeasts) was determined under the same fermentationconditions (10% initial glucose, 30 �C, and pH of 5.0) (Fig. 1A). Theresults showed that batch fermentation with yeast immobilized incalcium alginate beads required less fermentation time to convertglucose to ethanol than did cells in batch fermentation. Fermenta-tion time to consume the glucose by free cells was about 23 h,whereas immobilized cells with 2% and 2.5% calcium alginatebeads required 20 and 22 h, respectively. No physical breakagewas observed with the 2% and 2.5% calcium alginate beads duringfermentation. Higher sodium alginate concentrations resulted indelayed glucose consumption because the surfaces of the calciumalginate beads were expected to hold more strongly and be moredense at higher concentrations, thus making it more difficult forglucose to percolate into the cell pores. The results showed that

Fig. 1. Comparison of glucose consumption: yeast immobilized in 2% and 2.5%alginate beads vs. free yeast cells with 10% initial glucose concentration (A), 10% vs.15% initial glucose concentration(B), at 30 �C, pH 5.0.

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the 2% alginate concentration provided the best balance betweenporosity and strength.

Ethanol productivity of immobilized cells was equal to or lowerthan that of free cells. Roukas (1994) reported that both free andimmobilized S. cerevisiae cells produce the same maximum ethanolconcentration under the same fermentation conditions as nonster-ilized carob pod extract in a batch-fed culture. Singh et al. (1998)also concluded that the concentrations of ethanol produced in bothfree and immobilized batch systems (Ca-alginate) were relativelysimilar. Mariam et al. (2009) reported that the ethanol yield ob-tained by free cells was 7.3% higher than the yield obtained fromimmobilized cells. However, the more common finding is that un-der the same fermentation conditions, immobilized yeast cellsyield more ethanol than do free cells. Yu et al. (2007) reported thatethanol productivity of immobilized cells in sorghum bagasse was2.24 times higher than that of free cells. However, Holcberg andMargalith (1981) reported that the rate of ethanol production byentrapped cells with agar, j-carrageenan, alginate, and polyacryl-amide gels was higher than that with free cells. Norton et al.(1995) reported a significant increase in yeast resistance againstethanol for immobilized cells compared with free cells. The resultsreported by Roukas (1994) and Mariam et al. (2009) are differentfrom the results shown in Fig. 1A, but the results by Yu et al.(2007), Holcberg and Margalith (1981), and Norton et al. (1995)are similar to ours.

Gilson and Thomas (1995) and Najafpour et al. (2004) describedthe effect of alginate concentration on fermentation products andproperties of alginate beads. They showed that beads with lowalginate (<1.5%) were too soft and easily breakable, whereas2–5% (w/w) alginate beads suffered no damage. Soft beads alsohad problems such as overgrowth and expansion of bead diameterwhen grown in sugar solution. Fermentation with >3% alginateconcentrations leads to decreases in glucose consumption and eth-anol production rates, and cell activity decreases at 6% alginateconcentration. Gilson and Thomas (1995) reported that 5.1-mm-diameter beads of 2% (w/w) alginate suffered some breakage,whereas 1.3–3.7-mm beads suffered no breakage. In the presentstudy, fresh 2% calcium alginate beads with a diameter of3.8 mm under first-batch fermentation showed no breakage, butreused 2% calcium alginate beads in the fifth-batch fermentationshowed some breakage and splits.

3.2. Effect of initial glucose concentration

Fig. 1B shows that 10% and 15% glucose solutions took 24 and36 h of fermentation time, respectively. Thomas et al. (1996) re-ported that S. cerevisiae produced and tolerated high ethanol con-centrations under appropriate environmental and nutritionalconditions. Wendhausen et al. (2001) found that ethanol yield de-pended strongly on initial glucose concentration during batch fer-mentation of sugar cane syrup by immobilized S. cerevisiae. Wadaet al. (1980) also examined the effect of 10–25% glucose concentra-tions on the growth of cells and on ethanol-producing activity incells immobilized with j-carrageenan. They found that an increasein glucose concentration reduced the growth rate of cells and thenumber of living cells in the gel. They concluded that supplying10% glucose medium allowed for the largest number of cells inthe gel. Liu et al. (2009) examined ethanol yield with differentsugar and molasses concentrations (9%, 12%, and 15%) in a magnet-ically stabilized fluidized bed reactor under a continuous fermenta-tion process with immobilized S. cerevisiae cells. The ethanol yielddecreased as the sugar concentration and feed dilution increased.At a higher sugar concentration, a longer retention time wasrequired to achieve higher ethanol yield. Another report estimatedthat maximum production is obtainable at a 15% sugar level byboth free and immobilized cells under batch fermentation (Mariamet al., 2009). However, at lower glucose concentrations (2.5–5%),sugar consumption trends show similar results (Najafpour et al.,2004). A longer retention time is required for higher sugar concen-tration, and ethanol yield decreases as sugar concentrationincreases (Holcberg and Margalith, 1981; Najafpour et al., 2004).The decreased fermentation yields with higher sugar concentra-tions are due to osmotic effects. Roukas et al. (1991) reported thatreduced water activity combined with plasmolysis causes adecrease in the fermentation rate and ethanol production abovea critical substrate concentration. As mentioned earlier, in ourstudy, 10% glucose led to a shorter fermentation time than thatfor 15% glucose, which also included a longer retention time.

3.3. Effect of temperature

Four different temperatures (30, 32.5, 35, and 37.5 �C) weretested to determine the optimal temperature for batch fermenta-tion with yeast immobilized in 2% alginate beads at pH 5.0(Fig. 2). The results showed that an increase in temperature from30 to 37.5 �C had a fluctuating effect on glucose conversion time:19.5, 20.5, 19, and 23 h, respectively, were needed to reach 90%ethanol conversion. Temperature values from 30 to 37.5 �C, except27.5 �C, showed very similar retention times for glucose consump-tion, but 30 �C at the same fermentation conditions was the opti-mal temperature using immobilized yeast under batch

Fig. 2. Comparison of glucose consumption by temperature, ranging from 27.5 to37.5 �C, with a 10% initial glucose concentration and pH 5.0.

Fig. 3. pH change vs. unregulated fermentation (A); Comparison of glucoseconsumption by pH, ranging from 4.5 to 5.5, with a 10% initial glucose concentra-tion at 30 �C (B).

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fermentation. Notably, this low-temperature fermentation reducedenergy consumption.

The influence of fermentation temperature (from 15 to 35 �C)on a mixed-strain population was studied by Torija et al. (2003).After 4 days of initial fermentation, the cell population increasedat high temperatures compared with low temperatures. The usualgrowth curve, with a series of short lags and exponential, station-ary, and decline phases, was observed at 25 and 30 �C, whereas at35 �C, many of the yeast cells died. A similar result was found whenthe effect of incubation temperature on ethanol production wasexamined in free and immobilized yeast at 15% initial sugar con-centration and pH 4.5 (Mariam et al., 2009). They estimated etha-nol production at different incubation temperatures (25–40 �C)and found that 30 �C supported maximum ethanol production byfree yeast cells after 120 h of incubation. Although immobilizedcells showed lower ethanol yield and sugar consumption com-pared with free cells, the overall optimal temperature of fermenta-tion was 30 �C. Another study reported that the ethanol yieldincreased from 75.79% to 89.89% as fermentation temperature in-creased from 28 to 37 �C (Liu and Shen, 2008). The highest ethanolyield was observed at a fermentation temperature of 37 �C. Theoptimum temperature of free S. cerevisiae fermentation is usuallyabout 30 �C (Torija et al., 2003). Liu and Shen (2008) found thatthe optimum temperature of fermentation with immobilized yeastis higher than that with free yeast. As shown in Fig. 2 and 30 �C wasthe optimal fermentation temperature without noticeable reten-tion time.

3.4. Effects of pH

pH changes were measured in unregulated culture mediumduring fermentation (Fig. 3A). To find the pH effect on fermenta-tion, we studied fermentation at pH 4.5, 5.0, and 5.5 and comparedthe results with those from a natural (unregulated) pH (Fig. 3B).Fig. 3A shows that the pH values decreased to 3.85 from 6.4 underunregulated conditions. Fig. 3B shows that regulated conditions arebetter than unregulated in terms of fermentation time. Among reg-ulated samples, pH 5.0 was the best, with >90% of glucose con-sumed in 19.5 h.

Valli et al. (2005) reported that cells derived from differentgrowth phases behave significantly differently at different pH val-ues. By decreasing the external pH from 7.0 to 2.2, a progressivereduction in the intracellular pH from 7.1 to 5.1 was observed inexponentially grown cells. However, stationary cells maintained

constant intracellular pH at around 6.1 when the external pHwas in the range of 7.0–5.5, and an intracellular pH of 5.5 wasmaintained by stationary cells even with pH values decreasingfrom 5.0 to 2.2.

Kourkoutas et al. (2004) reported that reduced intracellular pHis attributable to increased permeability of the plasma membraneto protons, which leads to higher consumption of ATP, causing in-creased glycolytic activity and glucose uptake. Mariam et al. (2009)reported that the rate of ethanol production by yeast cells washighly affected by the pH of the fermentation medium. S. cerevisiaeshowed maximum growth under acidic conditions. The optimumpH for free-yeast ethanol fermentation in their study was pH4.8–5.0, whereas Liu and Shen (2008) obtained a maximum etha-nol yield at pH 5.0 and suggested that the pores on the calciumalginate matrix are large enough to allow nutritional materialsand inorganic salt ions to pass freely from the outside of immobi-lized yeast particles to the inside. H+ concentration in the fermen-tation solution changed the charge quantities of the yeast plasmamembrane. As a result, the permeability of the yeast plasma mem-brane for some materials and ions including carbon sources, nitro-gen sources, and other inorganic salt ions changed with H+

concentration. Carmelo et al. (1997) showed that higher pH af-fected the activity of plasma membrane ATPase and the cytosolicand vacuolar pH of yeast cells. In this study, pH 5 was the best ina regulated system. This result was in good agreement with the re-port of Liu and Shen (2008).

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3.5. Ethanol production by fresh and reused beads

Glucose consumption and ethanol production were studied as afunction of time with free cells and reused beads at 30 �C, pH 5.0(Fig. 4). Reused beads provided overall better performance com-pared with free cells and fresh beads. Among the reused cases,the three-time reused beads produced the best results consideringboth fermentation time (10 h) and ethanol yield. Fermentationtimes with two, four, and five-time reused beads were also favor-able: 14 h compared to 24 h with fresh beads and 36 h for free cells(Fig. 4A). For ethanol production, fresh or reused beads producedclose to a theoretical ethanol yield of 100% (0.51 g ethanol/g glu-cose), but free cells produced a lower yield (88%) (Fig. 4B).

Mariam et al. (2009) reported that during six consecutive batchruns with immobilized yeast cells, ethanol yield increased up tothe third batch run, and maximum ethanol yield was obtained inthe fourth batch run. Additionally, both sugar consumption rateand ethanol yield decreased sharply in the fifth and sixth batchruns. The best ethanol yield from the fourth batch run reportedby Mariam et al. (2009) was approximately 86%, whereas all runswith reused cases were close to 100% in the present study. It is alsoimportant to note that their fermentation time was significantlylonger than ours, at an average of 120 h compared to 10–14 h.The reason for this marked difference in performance is that Mar-iam and colleagues ran their fermentation with a 15% initial glu-cose solution at pH 4.5 and 30 �C. A comparison of our resultswith those from the literature is shown in Table 1. This Tableshows that the previous fermentation time was >24 h, but our re-sults required 10–14 h. Therefore, ethanol productivity in the pres-ent study was about 1.7-fold higher than previous results.

Fig. 4. Comparison of glucose consumption vs. time (A) and ethanol production vs.time (B) among free cells, fresh 2% alginate beads, and 2% reused alginate beads at30 �C, pH 5.0.

3.6. Morphological characteristics and population of yeast cells onalginate beads

We observed differences in color and diameter between freshalginate beads and reused beads (data not shown). The color differ-ence between fresh white beads and reused yellow beads was pos-sibly caused by the change in yeast concentration between the twosamples (Fig. 5A and B). The alginate beads swelled after fermenta-tion; the average diameter increased from 3.8 mm for fresh beadsto 4.2 mm for reused beads. Thus, an average 10.5% increase inbead diameters occurred with an �180% bulk volume increase.The SEM images revealed that the population of yeast cells onthe surface of reused beads was significantly higher than that onthe surface of fresh beads (Fig. 5A and B). The population of yeastcells on the fresh alginate beads was scattered across the surface,

Table 1A comparison of the experimental condition and ethanol production.

Lee et al. (This work) Kiran et al.

Strain S. cerevisiae (KCTC 7906) S. cerevisiaeTemperature (�C) 30 30Beads 2% Ca-alginate (Ø 3.8 mm,

430ea beads/L, Air-dried 9.6 g,)3% Ca-algin250ea bead

Required fermentation times (h) 10–14 48 (about 9ethanol wa24 h)

Substrate concentration 10% Glucose 20% Glucos

Maximum amount of ethanol (g/L)/1st batch (g/L, fermentation times)

51.1/(51.1, 24 h) 92/(36, 48

Ethanol yield (%) 100 90.1Repeated cycle (Maximum yield

batch)5 (3) 2–5 Batch remainedalmost constant

6 (6)

pH 5.0 5.5Volumetric ethanol productivity (g/

L h)5.11 1.94

except for some agglutinated yeast cells in the middle (Fig. 5A),whereas the entire surface of reused beads was densely coveredwith yeast cells (Fig. 5B).

The TEM images also showed that the population of yeast cellswithin a reused bead was significantly higher than that in freshbeads (Fig. 5C and D). Only one yeast cell per 1740 lm2 yeast cellswas found on fresh beads, but reused beads showed about seven

(2000) Mariam et al. (2009) Chandel et al. (2009)

(VS3) S. cerevisiae (GC-IIB31) S. cerevisiae (VS3)30 30

ate (Ø 5 mm,s/L)

2% Ca-alginate (2 g yeastcells)

Ten pre-weighed dry wildsugarcane stalks

0% of thes produced in

24 36

e 15% Sugar from canemolasses using 0.5% sulfuricacid

5.4% Sugar from S.spontaneum enzymatichydrolysate

h) 75.6/(53.8, 24 h) 22.85/(21.6, 36 h)

100.6 906 (4) Sharply in the 5th and6th batches

13 (2–8) 2–8 Batch remainedalmost constant

4.5 5.53.15 0.63

Fig. 5. Comparison of ultrastructural characteristics between fresh beads and reused beads. Scanning electron microscopy images of the fresh bead surface beforefermentation (A) and a reused bead after the second fermentation (B). Transmission electron microscopy images from yeast cells with fresh beads before fermentation (C) andreused beads after the second fermentation (D). Bars: A and B = 2 lm, C and D = 500 nm.

Fig. 6. Ultrastructural characteristics of fresh calcium alginate beads before fermentation, calcium alginate beads after the first fermentation, and reused calcium alginatebeads after the third fermentation. The alginate matrix model known as the ‘‘egg-box’’ (A), and transmission electron microscopy images of calcium alginate matrix structuresof fresh beads before fermentation (B), calcium alginate beads after first fermentation (C), reused calcium alginate beads after the third fermentation (D). Bars: 200 nm.

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K.H. Lee et al. / Bioresource Technology 102 (2011) 8191–8198 8197

yeast cells in the same area. Christensen et al. (1990) suggested thealginate matrix structure known as the ‘‘egg-box’’ model (Fig. 6A).We first identified the yeast cell suspension in the calcium alginateand the egg-box structures of the alginate matrix (Fig. 6). It is verydifficult to observe calcium alginate beads with an electron micro-scope because they shrink during dehydration for electron-micro-scope pretreatment, so the beads do not maintain their originalmorphological characteristics. This problem was solved using amodified pretreatment method. Fig. 6 shows the ultrastructuralcharacteristics of fresh calcium alginate beads before fermentationand after the first fermentation, and reused calcium alginate beadafter the third fermentation. The sizes of the micropores known asthe ‘‘egg-box’’ were measured as 177.9 ± 22.9 nm in fresh calciumalginate beads before fermentation (Fig. 6B), 177.2 ± 82.8 nm incalcium alginate beads after the first fermentation (Fig. 6C), and194.4 ± 27.4 nm in reused calcium alginate beads after the thirdfermentation (Fig. 6D). The size of micropores composed of micro-chains of alginate polymers increased under repeated batch fer-mentation, which may have increased bead diameters and thebulk volume.

The increased volume, diameter, and micropore sizes of reusedbeads caused by yeast population growth and water swelling mostlikely enhances the permeability of glucose from the surface in thereused beads to the pores, where yeast cells are located. This inturn caused an increase in the glucose conversion rates to ethanol,thereby reducing fermentation time. Gilson and Thomas (1995)studied the effect of different bead diameters of several freshlyprepared beads on glucose consumption and ethanol productionrates. Their results showed that glucose consumption and ethanolyield decreased with increasing diameter of fresh beads. However,in this study with reused beads in which the diameter increased,we showed an increased rate of ethanol production with reducedfermentation time. Gilson and Thomas (1995) compared freshbeads of different diameters, whereas we compared fresh beadsto reused beads, and the diameter increased with fermentation.Our results imply that the porosity and high population of yeastcells in the beads, and not the physical size of beads, are importantfactors for the rapid conversion of glucose to ethanol with a highethanol yield. We identified, for the first time, the distribution ofyeast cells in a calcium alginate and alginate matrix structuretermed the ‘‘egg-box’’ model by Christensen et al. (1990).

4. Conclusions

Results with all immobilized yeast produced 100% ethanolyields, whereas the free cells yielded 88%. Ethanol production inthis study was about 1.7-fold higher than that reported in previousstudies. A 100% ethanol yield using the immobilized batch processand a fast fermentation time of 10–14 h with no or fewer metabolicby-products and no residual starting material such as glucose inthe final product solution is the most favorable fermentation pro-cesses for economical bioethanol production.

Acknowledgements

This work was supported by Priority Research CentersProgram (Project No. 2011-0018394) and World Class Universityproject (R31-2009-000-20025-0) through the National ResearchFoundation of Korea (NRF) funded by the Ministry of Education,Science and Technology, and by the New & Renewable Energy ofthe Korea Institute of Energy Technology Evaluating Planning(2010T100100573) grant funded by the Korea Government Ministryof Knowledge Economy to H-J. Bae. ISC and YGK are grateful for theBK21 program provided by the Ministry of Education.

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