Exopolysaccharide production by a genetically engineered Enterobacter cloacae strain for microbial enhanced oil recovery

Bioresource Technology 102 (2011) 6153–6158

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Exopolysaccharide production by a genetically engineered Enterobacter cloacae strain for microbial enhanced oil recovery Shanshan Sun a, Zhongzhi Zhang a,⇑, Yijing Luo a, Weizhang Zhong a, Meng Xiao a, Wenjing Yi a, Li Yu b, Pengcheng Fu a a b

State Key Laboratory of Heavy Oil Processing, Faculty of Chemical Engineering, China University of Petroleum, Beijing 102249, PR China Institute of Porous Flow & Fluid Mechanics, Research Institute of Petroleum Exploration & Development, Langfang, Hebei Province 065007, PR China

a r t i c l e

i n f o

Article history: Received 10 November 2010 Received in revised form 1 March 2011 Accepted 2 March 2011 Available online 9 March 2011 Keywords: Microbial enhanced oil recovery (MEOR) Water-insoluble exopolysaccharide Electrotransformation Genomic DNA Core flooding

a b s t r a c t Microbial enhanced oil recovery (MEOR) is a petroleum biotechnology for manipulating function and/or structure of microbial environments existing in oil reservoirs for prolonged exploitation of the largest source of energy. In this study, an Enterobacter cloacae which is capable of producing water-insoluble biopolymers at 37 °C and a thermophilic Geobacillus strain were used to construct an engineered strain for exopolysaccharide production at higher temperature. The resultant transformants, GW3-3.0, could produce exopolysaccharide up to 8.83 g l1 in molasses medium at 54 °C. This elevated temperature was within the same temperature range as that for many oil reservoirs. The transformants had stable genetic phenotype which was genetically fingerprinted by RAPD analysis. Core flooding experiments were carried out to ensure effective controlled profile for the simulation of oil recovery. The results have demonstrated that this approach has a promising application potential in MEOR. Ó 2011 Elsevier Ltd. All rights reserved.

1. Introduction Biotechnology has found an increasing application in petroleum engineering. Microbial enhanced oil recovery (MEOR) is one of such biological efforts which may impact the petroleum industry substantially in current energy shortage period. For the crude oil production, it was concluded that the recovery percentage by both primary and secondary methods is typically 30–50% (Brown, 2010; Tzimas et al., 2005). Current tertiary recovery, or enhanced oil recovery (EOR), allows another 5–15% of the reservoir’s residual oil to be recovered (Green and Willhite, 1998). Therefore, it is important to develop novel approaches to improve the efficiency of EOR of oil entrapped in porous media to increase economic profits. MEOR is a petroleum biotechnology for manipulating function and/or structure of microbial environments existing in oil reservoirs for prolonged exploitation of the largest source of energy. MEOR possesses several advantages over other enhanced oil recoveries, which include low costs, broad applications, stable bacterial activity, environmentally friendly, and so on (Lazar et al., 2007). The microorganisms used in MEOR may produce a variety of bioproducts, such as biosurfactants, biopolymers, biomass, acids, solvents, gases and enzymes which could be utilized to extend the life of the oil reservoirs (Sen, 2008).

⇑ Corresponding author. Tel.: +86 10 89734284; fax: +86 10 89733974. E-mail address: [email protected] (Z. Zhang). 0960-8524/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2011.03.005

One of the main challenges in the oil production by water flooding is the variation of reservoir permeability. This can be circumvented by selective plugging of high permeability areas (Nemati et al., 2005). The role of microbial biopolymer in enhanced oil recovery is to improve the volumetric sweeping efficiency of waterflood by selective plugging of high permeability zones or water-invaded zones (Lazar et al., 2007; Yakimov et al., 1997). Several bacteria have been found to be capable of secreting biopolymers. For instance, xanthan gum can be produced by Xanthomonas campestris (Becker et al., 1998), glucan is found to be secreted by Lactobacillus suebicus, Pediococcus parvulus or P. parvulus (Garai-Ibabe et al., 2010), dextran is produced by Leuconostoc mesenteroides (Kim and Fogler, 1999), lewan is produced by Halomonas species (Poli et al., 2009) or Bacillus licheniformis (Garai-Ibabe et al., 2010; Liu et al., 2010; Ramsay et al., 1989) and pullulan is one of the metabolic products by Aureobasidium pullulans (Singh et al., 2009). Enterobacter cloacae is a gram-negative bacteria which is capable of producing insoluble polymer. This type of microbe can be grown at 4–60 °C and pH 5–7 (Prasertsan et al., 2006). It is reported that the yield of exopolysaccharide from E. cloacae can be up to 7.28 g l1 at optimum environmental conditions and its specific growth rate is 3 times higher than that of pullulan production from A. pullulan (Prasertsan et al., 2008). An E. cloacae strain has been identified which could produce water-insoluble extracellular polysaccharide at 37 °C when glucose was supplied. The E. cloacae was named JD and will be used in this study. The exopolysaccharide produced by JD could be used for

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sealing high permeability areas in the mature oil reservoirs. The aim of this paper was to construct an engineered E. cloacae strain from JD and a thermophilic Geobacillus strain by electrotransformation. The transformants could produce exopolysaccharide at elevated temperature (up to 50 °C). The engineered E. cloacae strain was characterized in liquid culture with growth parameters such as pH, temperature and salt tolerance in order to seek for optimal conditions for cell growth and exopolysaccharide production, Random amplification of polymorphic DNA (RAPD) PCR was used to confirm the success of DNA transformation. Core flooding experiments were conducted to evaluate the effects of simulated MEOR. 2. Methods 2.1. Microorganisms The E. cloacae strain, JD, was originally isolated from the wastewater in Jilin Oil Field of PetroChina Company Ltd., Jilin province, China. This bacterium was, ampicillin resistant and exopolysaccharide-producing. However, when the growth temperature was higher than 37 °C, JD would cease to produce exopolysaccharide. The Geobacillus sp. bacteria, GW3 and GW4, were isolated from the wastewater in Daqing Oil Field of PetroChina Company Ltd., Heilongjiang province, China. The two donor strains used in this work, GW3 and GW4, were gram-positive bacteria, ampicillin sensitive, and their most suitable growth temperature was in the range of 60–70 °C. GW3 and GW4 were not capable of producing biopolymer under any condition.

v/v/v) and centrifuged at 14,000 rpm for 5 min. DNA was precipitated from supernatant by 0.6 volume of isopropanol, washed with 1 ml 70% ice-cold ethanol, dissolved in 50 ll sterile H2O containing 20 lg RNase (Takara) per milliliter, and stored at 20 °C. The DNA was semiquantified in 0.8% agarose gel in 1  Tris–Acetic–EDTA and visualized by staining with ethidium bromide (EB). The purity and concentration of DNA were estimated by spectrophotometry (Zhang et al., 2003). 2.4. Electrotransformation Eighty microliter of competent JD cells from 70 °C refrigerator was thawed in ice-water mixture, mixed with 1 ll GW3 or GW4 genomic DNA (1 lg ll1) and incubated on ice for 5 min. Then the mixture was transferred to a chilled 2 mm electroporation cuvette (Bio-Rad, California, USA). The samples were electroporated in the MicroPulser™ Electroporation (Bio-Rad) at voltages of 2.8 kV, 2.9 kV and 3.0 kV, respectively. After the electric pulse, 400 ll of SOC medium (2% tryptone [m/v], 0.5% yeast extract [m/v], 10 mM NaCl, 2.5 mM KCl, 10 Mm MgCl2, 10 mM MgSO4, 20 mM glucose) was added into the cuvette immediately. The cell suspension was transferred to a 1.5 ml Eppendorf tube and incubated at 37 °C for 1 h with shaking at 110 rpm. Finally, all of cell culture was plated on 100 lg ml1 ampicillin selective complete medium (10 g of tryptone, 5 g of yeast extract, 10 g of glucose, 5 g of beef extract and 5 g of sodium chloride per liter) and incubated at 50 °C overnight to select transformants. Positive clones were further verified with liquid ampicillin selective complete medium in static culture at 50 °C.

2.2. Preparation of electrocompetent cells

2.5. RAPD-PCR assay

Competent JD cells for electrotransformation were prepared with the protocol as follows (Dorella et al., 2006; Olubajo and Bacon, 2008): 5 ml of Luria–Bertrani (LB) broth (10 g of tryptone, 5 g of yeast extract, and 10 g of sodium chloride per liter) in a test tube was inoculated with a single colony of JD and incubated at 37 °C with shaking at 200 rpm overnight. Then the overnight culture was diluted 10-fold with 45 ml LB broth and incubated with shaking at 250 rpm at 37 °C to an optical density of OD600 nm = 0.6–1.0. The culture was chilled on ice for 30 min; 1 ml of JD cell culture broth was sampled and then harvested by centrifugation at 13000 rpm at 4 °C for 30 s. The supernatant was removed, and the cell pellets were resuspended with 1 ml of icecold 10% glycerol in water (v/v), and centrifuged at 13000 rpm for 30 s at 4 °C. This step was repeated three times. Following the last centrifugation, the cells were resuspended in 400 ll of 10% ice-cold glycerol and stored at 70 °C.

Random amplification of polymorphic DNA (RAPD) is one of PCR-based DNA fingerprinting methods (Dubey et al., 2006). It uses short primers of arbitrary nucleotide sequence to reproducibly amplify segments of genomic DNA to detect bases changes (Williams et al., 1990). The genomic DNA of transformants and their parents was extracted as mentioned above. Twentyfive microliter volume of PCR reaction mixtures contained 2.5 ll of a 10-fold PCR buffer without MgCl2, 1.5 ll of 25 mM of MgCl2, 2 ll of 2.5 mM dNTP (Promega, Wisconsin, USA), 1 ll of 5 lM of primer (Sangon, Shanghai, China; Table 1), 100 ng of template DNA, 1.5 ll of 5 u ll1 DNA polymerase (Promega), and purified water to the final volume. The amplifications were carried out in a DNA thermal cycler Techgene (Techne, Cambridge, UK) using the PCR protocol including an initial denaturation at 94 °C for 4 min, 45 cycles of 94 °C for 1 min, 36 °C for 1 min, 72 °C for 2 min and final extension at 72 °C for 5 min. The RAPD-PCR products were separated in 0.5  Tris–Borate–EDTA gel containing 2% agarose and stained with ethidium bromide. The gels were photographed with a gel imaging system (Vilber Lourmat, Marne la Vallée, France).

2.3. Extraction of GW genomic DNA GW genomic DNA was extracted according to a cetryltrimethylammonium bromide (CTAB)-NaCl protocol (Conn and Franco, 2004), with some modifications. Briefly, 1 ml of GW3/GW4 cell culture broth was sampled and harvested by centrifugation at 13000 rpm at 4 °C for 30 s. The supernatant was removed, and resuspended with 567 ll TE buffer (10 mM Tris–HCl pH 8.0, 1 mM EDTA) and 3 ll 20 mg ml1 proteinase K (Takara, Shiga, Japan). This suspension was lysed with 30 ll 10% sodium dodecyl sulfate (SDS) at 37 °C for 1 h. Subsequently, 100 ll of 5 M NaCl and 80 ll of CTAB-NaCl (700 mM NaCl, 275 mM CTAB) were added, and the mixture was incubated at 65 °C for 10 min. The cell lysates were extracted first with 780 ll of chloroform-isoamyl alcohol (24:1, v/v) at room temperature for 1 min with slow shaking. After centrifugation (10,000 rpm, 5 min), the supernatant was mixed with one volume of phenol–chloroform-isoamyl alcohol (25:24:1,

Table 1 DNA sequence of random primers used for RAPD-PCR. Primer

DNA sequence (50 –30 )

Primer

DNA sequence(50 –30 )

S12 S86 S208 S265 S366 S381 S501 S511 S1142 S1149

CCTTGACGCA GTGCCTAACC AACGGCGACA GGCGGATAAG CACCTTTCCC GGCATGACCT TGCGGGTCCT GTAGCCGTCT AATCCGCTGG CCAGATGGGG

S1293 S1343 S1366 S1467 S1480 S2006 S2007 S2008 S2154 S2160

CTGACTTCCC TTTCCGGGAG CCTTCGGAGG GTGTCAGTGG TTGACCCCAG GGACGACCGT GGGTCGCATC CCACAGCCGA ACCGTGGGTG CACCGACATC

S. Sun et al. / Bioresource Technology 102 (2011) 6153–6158

3.2. Construction of engineered bacteria

Table 2 Composition of model brinea used in core flooding experiment. Constituent

K++ Na+

Ca2+

Mg2+

Cl

SO42

HCO3

Total salinity

Concentration (mg l1)

3945

159

73

6507

19

96

10799

a

Water type was CaCl2, the viscosity of the brine at 50 °C was 1 mPa s, pH was

7.5.

2.6. Core flooding experiment The effect of exopolysaccharide produced by transformants applied to selective plugging of high permeability zones was evaluated by core flooding experiment. The process was conducted at 50 °C to simulate the actual temperature of the oil reservoirs at Fang 19 Unit, Dagang Oil Field of PetroChina, Tianjin, China. The artificial core used in this study was cylinder of 71.2 mm in length and 25 mm in diameter. The artificial core was saturated with sterile brine (Table 2) after vacuum pumping. Then the saturated core was inserted into metal core holders and flooded with sterile brine until the pressure became stable. After first water flooding, two porous volume (PV) of bacteria solution (109 cells ml1) mixed with complete medium (1:25, v/v) were injected into the cores and incubated for 3 days at 50 °C. In the period of subsequent water flooding, the cores were flooded with the same brine. The flow rate for the core flooding was set to be 1 ml min1. The pressure was recorded to determine the effective permeabilities for model brine before (Kwb) and after bacterial treatment (Kwa), and the permeabilities were calculated using Darcy’s Law. The blocking ratio (R) were calculated by the formulas as follows (Zhu et al., 2009):

R ¼ ð1 

K wa Þ  100% K wb

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ð1Þ

3. Results and discussion 3.1. Purity of genomic DNA extracted from donor strains Gel electrophoresis was used to estimate the quality of genomic DNA. The extracted DNA was integrated and had high molecular weight (Fig. 1). The concentration was assessed by absorbance of ultraviolet light at wavelengths of 260 nm, and the purity of DNA was determined by the ratio OD260/OD280. The concentration of GW3 and GW4 genomic DNA was 1.05 and 1.08 lg ll1, respectively. OD260/OD280 ratio of GW3 and GW4 was 1.87 and 1.85 which demonstrated the extracted DNA was of high purity (Zhang et al., 2003).

Fig. 1. Electrophoretic analysis of genomic DNA in 0.8% agarose gel. Lane 1, GW3; lane 2, GW4; lane 3, 1 kb DNA marker.

Transformants following electroporation were selected on complete medium plate with ampicillin and cultured at 50 °C. Positive clone was then transferred to 2 ml working volume of complete medium containing ampicillin in static culture at 50 °C for 24 h to check if the transformant produced water-insoluble extracellular polysaccharide. As a result, five transformants named GW32.8, GW3-2.9, GW3-3.0, GW4-2.8 and GW4-3.0 were confirmed which could produce exopolysaccharide at 50 °C. The donor strain of GW3-2.8, GW3-2.9 and GW3-3.0 was GW3, and that of GW4-2.8 and GW4-3.0 was GW4. The numbers following the hyphen represented the voltages used in electroporation. It has been found that the electrotransformation efficiency relied on activity of the electrocompetent JD cells, harvested genomic DNA from GW3/GW4 and the electroporation media. In order to enhance the transformation efficiencies, genomic DNA transformed into competent JD cells is preferred to be in circular plasmids, based on the studies in the literature (Chaurasia et al., 2008; Dorella et al., 2006). On the other hand, although linear plasmids are about 103–104 fold less efficient than circular plasmids (Shigekawa and Dower, 1989; Simon and Ferretti, 1991), linear plasmids have shown to be more efficiently in integration into the host genome according to MicroPulser™ Electroporation Apparatus Operating Instructions and Applications Guide. In this study, the fragmented genomic DNA from thermophilic GW3 or GW4 cells was transformed into the JD cells and the new phenotype (exopolysaccharide production at elevated temperature of 50 °C) of transformants showed no changes after 20 subcultures (data not shown). It is thus obvious that the heterologous DNA has been integrated into the host chromosome and the DNA transformation was stable. Exopolysaccharide yield was determined by the method described by Prasertsan et al. (2008). The productivity of transformants was evaluated in the form of the dry weight of exopolysaccharide. The results indicated that all of the constructs, except GW3-2.9, could produce more biopolymer at 50 °C than that by JD cells cultured at 37 °C. Among them, GW3-3.0 and GW4-2.8 could produce more exopolysaccharide at the same condition than other transformants were chosen for the further experiments. 3.3. Characteristics of transformants The effects of temperature, salt concentration and pH on the performance of exopolysaccharide producing transformants were evaluated. (Fig. 2). It needs to be noted that so far all microorganisms in bioindustry for commercial polysaccharide-production are mesophiles (Prasertsan et al., 2008). For instance, the optimal temperature for polysaccharide formation by Aureobasidium pullulans was 24 °C (McNeil and Kristiansen, 1990). The optimal temperature for E. cloacae to produce exopolysaccharide was 30 °C (Prasertsan et al., 2008). In comparison, the transformants derived from this study have shown to produce more exopolysaccharide at higher temperatures. In Fig. 2a, it can be seen that the biopolymer produced by GW3-3.0 at 54 °C was the highest. In the same figure, it is found that the optimal temperature for GW4-2.8 to produce exopolysaccharide was 51 °C. Furthermore, the exopolysaccharide yield by GW3-3.0 at 54 °C (8.83 g l1) was similar to that by B. licheniformis at 30 °C (9.02 g l1) (Liu et al., 2010). This study indicates that further increase in temperature would result in inhibition to the engineered strains. For example, when the growth temperature was set to be 59 °C, the transformants were significantly inhibited and could produce almost no polysaccharide. Fig. 2b shows that the transformants tolerate higher salt concentrations than the wild type JD strain. The data is unexpectedly

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b

10

GW3-3.0 GW4-2.8

Dry weight of exopolysaccharide (g/l)

9

Dry weight of exopolysaccharide (g/l)

a

8 7 6 5 4 3 2 1 0

7 GW3-3.0 GW4-2.8 JD

6 5 4 3 2 1 0

46

47

48

49

50

51

52

53

54

55

56

57

58

59

60

15000

20000

o

Temperature ( C)

Weight of exopolysaccharide (g/l)

70

Wet weight of exopolysaccharide Dry weight of exopolysaccharide

100 Moisture content

90 60 50

80

40 70

30 20

60 10 50

0 JD

GW3-3.0

GW4-2.8

d

7

Dry weight of exopolysaccharide (g/l)

80

Moisture concent (%)

c

25000

30000

Salinity (mg/l)

6

GW4-2.8 GW3-3.0

5 4 3 2 1 0 4

5

6

7

8

9

10

pH Fig. 2. Characterizations of transformants producing expolysaccharide. (a) Effect of temperature on expolysaccharide production by GW3-3.0 and GW4-2.8 strains. Cultures were grown in molasses medium (40 ml molasses, 1 g (NH4)2HPO4, and 10 g NaCl per liter) at 47, 49, 51, 54, 56, and 59 °C. (b) Effect of NaCl concentration on transformants and JD producing expolysaccharide. GW3-3.0 and GW4-2.8 strains were cultured at 50 °C and JD at 37 °C in molasses medium with 15000, 20000, 25000 and 30000 mg l1 NaCl. (c) Moisture content of wet exopolysaccharide produced by JD, GW3-3.0 and GW4-2.8, respectively. (d) pH effects on the activity of GW4-2.8 and GW3-3.0 producing biopolymer at 50 °C. The pH of molasses medium was varied stepwise from 4.0 to 10.0 with 1.0 increment.

interesting that the exopolysaccharide production would be increased with the aid of salinity. It is assumed that salinity is in a kind of stress on the transformants, the cells increase the exopolysaccharide production to adapt themselves to the harsh environment. Fig. 2b also indicates that the wild type JD cells produced less polysaccharide than GW3-3.0 and GW4-2.8 when the medium contained molasses instead of glucose which suggested that the transformants were more efficient in the consumption of molasses than the JD cells. While the moisture content of wet exopolysaccharide produced by transformants (88%) was a little lower than that of JD (91%) (Fig. 2c). It is obvious that neutral pH was in favor of exopolysaccharide production by the transformants (see Fig. 2d). The transformant approximately produced more than 5 g l1 exopolysaccharide when the range of pH was between 6.0 and 9.0 (Fig. 2d). This is different from the biopolymer accumulation by some microorganisms. For example, Lactobacillus suebicus could produce glucan at pH 1.8 (Garai-Ibabe et al., 2010). The optimal yield of polysaccharide by GW3-3.0 was obtained at pH 7.0–8.0, it was similar to the literature report for cell growth of E. cloacae (i.e. the optimal pH for E. cloacae WD7 was 7.0, (Prasertsan et al., 2008). The experiment results showed that GW3-3.0 or GW4-2.8 would produce very little exopolysaccharide at pH 3 or lower.

3.4. RAPD analysis RAPD technology is simple and cost effective for genetic fingerprinting (Tyagi et al., 2007). It was generally used to analyse the bacterial diversity (Miyatake and Iwabuchi, 2005) and to assess the genetic variations of tissue culture plants (Martins et al., 2004). RAPD has been reported to be used for the investigation of the genetic relationship between protoplast fusant and the parental strains (Zhang et al., 2009). RAPD-PCR assay was used in this work in order to reveal the genetic relationship between the transformants and their parent cells. It was observed that there were four primers, S2006 (50 GGACGACCGT-30 ), S2007 (50 -GGGTCGCATC -30 ), S2008 (50 -CCACAGCCGA-30 ), and S2154 (50 -ACCGTGGGTG-30 ) been useful. The fragments amplified by these four selected primers were reproducible and their size was mostly in the range of 0.25–3 kb. In total, GW3, GW3-3.0, JD, GW4-2.8 and GW4 produced 26, 23, 34, 20 and 23 bands with the primers, respectively. The genetic similarity between GW3-3.0 and its parent strains GW3 and JD was 4.35% and 39.13%, respectively. Transformant GW4-2.8 shared genetic similarity of 5.00%, and 45.00% with GW4 and JD, respectively (Fig. 3). The results indicated that the transformants obtained more genetic information from the receptor JD strain.

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12 11

Bacteria injection

10 9

Pressure (kPa)

8 7 6

Subsequent water flooding

5 4 3

Water flooding

2 1 0 0

2

4

6

8

10

12 14

16 18

20 22 24 26 28

Water injected (PV) Fig. 4. Pressure variation during water flooding, bacteria injection and subsequent water flooding.

Fig. 3. Amplification profiles of RAPD fingerprinting analysis using the primers of S2006, S2007, S2008 and S2154. Lane 1, GW3; lane 2, GW3-3.0; lane 3, JD; lane 4, GW4-2.8; lane 5, GW4; lane M, D2000 plus DNA Ladder.

There were genetic differences among different transformants as shown from the data in Fig. 3. The genetic difference between GW3-3.0 and GW4-2.8 was 6.98% (Fig. 3) from RAPD analysis, which provided a more rigorous genetic proof to illustrate the discrepancy of transformants. 3.5. Microbial selective plugging Core flood experiments have been extensively conducted to evaluate MEOR (Cusack et al., 1992; Davey et al., 1998; Joshi et al., 2008; Suthar et al., 2008). There are three types of cores, namely, natural reservoir core, artificial core and sand pack column used in physical simulation experiments. Artificial cores are much more economical than natural ones. The structure of artificial cores is more authentic than the sand pack columns when observed in scanning electron microscope (SEM). These facts were taken into account when artificial cores were chosen to evaluate the efficiency of selective plugging by transformant GW3-3.0. The pressure increased from 1.85 kPa to 2.8 kPa during the injection of microbial cell cultures. After 3 days of incubation at 50 °C, the pressure within the artificial cores was increased because of the growth of microbial cells and the production of exopolysaccharide in the high permeability zones. Then the pressure became stable at a maximum of 11.5 kPa at the end of subsequent water flooding (Fig. 4). The blocking ratio representing the reduction in permeability of artificial cores was 83.9% which demonstrated that GW3-3.0 could be used to redirect the injection water around the plugged areas and into low permeability zones where the oil was left. Nevertheless, the permeability reduction was lower than that reported by Davey et al. by injection of phosphate and maltodextrin to stimulate indigenous bacteria (>90%) (Davey et al., 1998). The SEM photograph of dried exopolysaccharide in a representative sample from the artificial core was another evidence in support of preferential plugging of the high permeability zones created by the secretion of exopolysaccharide from GW3-3.0. Since 88.3% of wet exopolysaccharide produced by GW3-3.0 was water (Fig. 2c), the size of exopolysaccharide observed in SEM was only 10% of the original one.

Although this work focused on the creation of engineered E. cloacae strains for exopolysaccharide production for MEOR purpose, the methodology developed here may be applicable to other microbial applications which require combination of existing cellular functions in the host cells with novel functions by introduction of heterologous genes or DNA fragment. 4. Conclusions In this paper, a bio-based MEOR approach within the context of petroleum biotechnology was developed. Using this protocol, the delivered DNA was the whole genomic DNA in place of amplifying the gene related heat-resistant. The fragment including the heatresistant genes instead of the whole genome was introduced into the receptor bacteria by electrotransformation and the engineered E. cloacae possessed a novel function in producing exopolysaccharide at elevated temperature. Although the mechanism was complex and unclear, the stable phenotype of transformants and RAPD analysis have demonstrated that the fragment related heat-resistant might be integrated into the genome of JD cells. Acknowledgements The authors are grateful to the State Key Laboratory of Heavy Oil of China for their technical support. The authors also appreciate very much the two anonymous reviewers for their critical and constructive comments that have improved this manuscript. References Becker, A., Katzen, F., Pühler, A., Ielpi, L., 1998. Xanthan gum biosynthesis and application: a biochemical/genetic perspective. Applied microbiology and Biotechnology 50 (2), 145–152. Brown, L., 2010. Microbial enhanced oil recovery (MEOR). Current Opinion in Microbiology 13 (3), 316–320. Chaurasia, A., Parasnis, A., Apte, S., 2008. An integrative expression vector for strain improvement and environmental applications of the nitrogen fixing cyanobacterium, Anabaena sp. strain PCC7120. Journal of microbiological methods 73 (2), 133–141. Conn, V., Franco, C., 2004. Analysis of the endophytic actinobacterial population in the roots of wheat (Triticum aestivum L.) by terminal restriction fragment length polymorphism and sequencing of 16S rRNA clones. Applied and Environmental Microbiology 70 (3), 1787. Cusack, F., Singh, S., Novosad, J., Chmilar, M., Blenkinsopp, S., Costerton, J., 1992. The use of ultramicrobacteria for selective plugging in oil recovery by waterflooding. International Meeting on Petroleum Engineering, Beijing, China, 24–27.

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S. Sun et al. / Bioresource Technology 102 (2011) 6153–6158

Davey, M., Gevertz, D., Wood, W., Clark, J., Jenneman, G., 1998. Microbial selective plugging of sandstone through stimulation of indigenous bacteria in a hypersaline oil reservoir. Geomicrobiology Journal 15 (4), 335–352. Dorella, F., Estevam, E., Cardoso, P., Savassi, B., Oliveira, S., Azevedo, V., Miyoshi, A., 2006. An improved protocol for electrotransformation of Corynebacterium pseudotuberculosis. Veterinary Microbiology 114 (3–4), 298–303. Dubey, S., Tripathi, A., Upadhyay, S., 2006. Exploration of soil bacterial communities for their potential as bioresource. Bioresource Technology 97 (17), 2217–2224. Garai-Ibabe, G., Due as, M., Irastorza, A., Sierra-Filardi, E., Werning, M., López, P., Corbí, A., Palencia, P., 2010. Naturally occurring 2-substituted (1, 3)-[beta]-Dglucan producing Lactobacillus suebicus and Pediococcus parvulus strains with potential utility in the production of functional foods. Bioresource technology 101 (23), 9254–9263. Green, D., Willhite, G., 1998. Enhanced Oil Recovery, SPE textbook series, volume 6. Society of Petroleum Engineers, Richardson, Texas. Joshi, S., Bharucha, C., Jha, S., Yadav, S., Nerurkar, A., Desai, A., 2008. Biosurfactant production using molasses and whey under thermophilic conditions. Bioresource technology 99 (1), 195–199. Kim, D., Fogler, H., 1999. The effects of exopolymers on cell morphology and culturability of Leuconostoc mesenteroides during starvation. Applied microbiology and Biotechnology 52 (6), 839–844. Lazar, I., Petrisor, I., Yen, T., 2007. Microbial enhanced oil recovery (MEOR). Petroleum Science and Technology 25 (11), 1353–1366. Liu, C., Lu, J., Lu, L., Liu, Y., Wang, F., Xiao, M., 2010. Isolation, structural characterization and immunological activity of an exopolysaccharide produced by Bacillus licheniformis 8–37-0–1. Bioresource technology 101 (14), 5528–5533. Martins, M., Sarmento, D., Oliveira, M., 2004. Genetic stability of micropropagated almond plantlets, as assessed by RAPD and ISSR markers. Plant Cell Reports 23 (7), 492–496. McNeil, B., Kristiansen, B., 1990. Temperature effects on polysaccharide formation by Aureobasidium pullulans in stirred tanks. Enzyme and Microbial Technology 12 (7), 521–526. Miyatake, F., Iwabuchi, K., 2005. Effect of high compost temperature on enzymatic activity and species diversity of culturable bacteria in cattle manure compost. Bioresource Technology 96 (16), 1821–1825. Nemati, M., Greene, E., Voordouw, G., 2005. Permeability profile modification using bacterially formed calcium carbonate: comparison with enzymic option. Process Biochemistry 40 (2), 925–933. Olubajo, B., Bacon, C., 2008. Electrotransformation of Bacillus mojavensis with fluorescent protein markers. Journal of Microbiological Methods 74 (2–3), 102– 105. Poli, A., Kazak, H., Gürleyendag, B., Tommonaro, G., Pieretti, G., ner, E., Nicolaus, B., 2009. High level synthesis of levan by a novel Halomonas species growing on defined media. Carbohydrate Polymers 78 (4), 651–657.

Prasertsan, P., Dermlim, W., Doelle, H., Kennedy, J., 2006. Screening, characterization and flocculating property of carbohydrate polymer from newly isolated Enterobacter cloacae WD7. Carbohydrate Polymers 66 (3), 289–297. Prasertsan, P., Wichienchot, S., Doelle, H., Kennedy, J., 2008. Optimization for biopolymer production by Enterobacter cloacae WD7. Carbohydrate Polymers 71 (3), 468–475. Ramsay, J., Cooper, D., Neufeld, R., 1989. Effects of oil reservoir conditions on the production of water-insoluble levan by Bacillus licheniformis. Geomicrobiology Journal 7 (3), 155–165. Sen, R., 2008. Biotechnology in petroleum recovery: The microbial EOR. Progress in Energy and Combustion Science 34 (6), 714–724. Shigekawa, K., Dower, W., 1989. Electroporation: a general approach to the introduction of macromolecules into prokaryotic and eukaryotic cells. Australian Journal of Biotechnology 3 (1), 56. Simon, D., Ferretti, J., 1991. Electrotransformation of Streptococcus pyogenes with plasmid and linear DNA. FEMS Microbiology Letters 82 (2), 219–224. Singh, R., Saini, G., Kennedy, J., 2009. Downstream processing and characterization of pullulan from a novel colour variant strain of Aureobasidium pullulans FB-1. Carbohydrate Polymers 78 (1), 89–94. Suthar, H., Hingurao, K., Desai, A., Nerurkar, A., 2008. Evaluation of bioemulsifier mediated Microbial Enhanced Oil Recovery using sand pack column. Journal of Microbiological Methods 75 (2), 225–230. Tyagi, R., Agrawal, A., Mahalakshmi, C., Hussain, Z., Tyagi, H., 2007. Low-cost media for in vitro conservation of turmeric (Curcuma longa L.) and genetic stability assessment using RAPD markers. In Vitro Cellular & Developmental BiologyPlant 43 (1), 51–58. Tzimas, E., Georgakaki, A., Cortes, C., Peteves, S. 2005. Enhanced Oil Recovery using Carbon Dioxide in the European Energy System. European Commission Joint Research Centre, Report EVR, 21985. Williams, J., Kubelik, A., Livak, K., Rafalski, J., Tingey, S., 1990. DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Research 18 (22), 6531. Yakimov, M., Amro, M., Bock, M., Boseker, K., Fredrickson, H., Kessel, D., Timmis, K., 1997. The potential of Bacillus licheniformis strains for in situ enhanced oil recovery. Journal of Petroleum Science and Engineering 18 (1–2), 147–160. Zhang, P., Wu, D., Ge, J., Zhu, Z., Feng, G., Yue, T., Lin, J., Zheng, H., 2003. Experimental inhibition of corneal neovascularization by endostatin gene transfection in vivo. Chinese Medical Journal 116 (12), 1869–1874. Zhang, X., Jia, H., Wu, B., Zhao, D., Li, W., Cheng, S., 2009. Genetic analysis of protoplast fusant Xhhh constructed for pharmaceutical wastewater treatment. Bioresource Technology 100 (6), 1910–1914. Zhu, K., Lu, S., Xu, T., Xing, W., Jiang, W., Wang, C., 2009. Laboratory tests of factors effecting core sample blocking ratio and flow back break down ratio. Petroleum Drilling Techniques 37 (4), 61–64.