A novel bioemulsifier from Geobacillus stearothermophilus A-2 and its potential application in microbial enhanced oil recovery

Abstract

Biosurfactants can improve the mobility of oils in porous media by changing the rock wettability and emulsifying oils, thus increasing the efficiency of crude oil recovery in the petroleum industry. Therefore, surfactant-producers play important roles in the microbial enhanced oil recovery (MEOR) process. In this study, a thermophilic, facultative anaerobic emulsifier-producing strain was isolated. The emulsifier produced and its potential applications in MEOR were investigated in the laboratory. The stain was identified as Geobacillus stearothermophilus, and was designated as A-2, which could use sodium acetate, which is relative abundant in reservoirs, as the carbon source. The produced bioemulsifier is a novel glycoprotein emulsifier, containing 71.4% sugar and 27.8% protein, wherein the monosaccharides were identified as mannose (33.5%), glucose (30.9%), galactose (29.7%), and glucuronic acid (5.9%); and the protein contained 17 types of amino acids. The bioemulsifier successfully emulsified various hydrocarbons at a wide range of salinity, temperature, and pH. Notably, the emulsion layer of diesel remained stable for 12 months at room temperature, with little change at the micron level in the particle size of oil droplets. Core flooding tests indicated that the fermentation broth of strain A-2 increased the oil recovery efficiency by 6.8% under lower oil saturation condition, showing potential applications in oil exploration in high-temperature oil reservoirs.

---

Introduction

Compared with the chemical flooding techniques, microbial enhanced oil recovery (MEOR) has been a rapidly growing technique in recent years because of its simple operation, cost-effectiveness and environmental friendliness.^1–4 MEOR utilizes microorganisms and their metabolites such as biosurfactants, polymers, and gases to recover residual oil underground.^5–8 Among these metabolites, biosurfactants can significantly improve the mobility of oils in porous media through changing rock wettability and emulsifying oil, and have attracted much attention worldwide. Thus, increasing studies are striving to isolate surfactant-producers, and attempt to use these isolated microorganisms to enhance oil recovery in high water-content reservoirs.^9,10 Petroleum reservoirs are barren ecosystems, which are generally unsuitable for microbial growth and surfactant production due to the lack of suitable nutrients and enough oxygen, and to high temperature constraints.^3,11,12 Therefore, it is still a challenge to select appropriate strains for the MEOR technique. Actually, organic acids can be readily detected and considered as important carbon sources for microbial survival, wherein acetate ions are the most abundant in oil reservoirs.¹³ It is certainly a good choice for MEOR to select a facultative anaerobe, which can efficiently use organic acids as carbon sources to produce oil-displacing agents, such as biosurfactants.

Biosurfactants are amphiphilic substances consisting of hydrophobic and hydrophilic groups.^14,15 Based on the molecular size, biosurfactants are divided into two groups: (i) low-molecular-weight biosurfactants that reduce the surface and interfacial tensions and (ii) high-molecular-weight biosurfactants, known as bioemulsifiers that are more effective in stabilizing oil–water emulsions without significantly reducing the surface tension.¹⁶ Compared with chemical surfactants, biosurfactants have low toxicity, high biocompatibility, and effectiveness stability in a wide range of temperatures, pH, and salinity.¹⁷ These advantages make them excellent emulsifiers, and foaming and dispersing agents. The effectiveness of biosurfactants in MEOR has been demonstrated by a large number of laboratory experiments and field trials that were conducted by introducing exogenous biosurfactant-producing bacteria or biosurfactants, and stimulating endogenous microbial to produce biosurfactants.^14,18–21

Geobacillus is a thermophilic and aerobic or facultative anaerobic bacillus,²² and was detected in oil reservoirs. The members of this genus can grow in a range of 40–70 °C. Some species can produce surfactants. G. pallidus XS2 and XS3 were isolated from oil contaminated soil samples and were described as hydrocarbon-degraders and surfactant-producers.²³ Jara et al. showed that G. stearothermophilus UCP968 could use agro-industrial substrates such as corn steep liquor and palm oil as substrates to produce biosurfactants.²⁴ However, little information is available on surfactants' structure, and even chemical composition. Moreover, our knowledge about the surfactants production by Geobacillus sp. when grown on different carbon sources is also very limited.

In this study, the produced emulsifier and potential application of a thermophilic, facultative anaerobe—G. stearothermophilus A-2 were investigated. The physicochemical properties of the bioemulsifier were characterized by spectroscopic, chromatographic, and physicochemical methods. The potential application of the strain in MEOR was evaluated through core-flooding tests.

Materials and methods

Isolation of thermophilic and facultative anaerobic surfactant-producers

The produced water was separated from an oil production well of Dagang petroleum reservoir with an average temperature of 73 °C. To enrich surfactants producers, a total of 10 mL produced water was inoculated into flasks with 100 mL minimal salt medium, which contained 0.3 g L⁻¹ KH₂PO₄, 1.25 g L⁻¹ Na₂HPO₄·12H₂O, 2.0 g L⁻¹ NH₄Cl, 0.2 g L⁻¹ MgSO₄·7H₂O, 0.01 g L⁻¹ CaCl₂·2H₂O, 0.36 g L⁻¹ FeSO₄·7H₂O, 0.5 g L⁻¹ yeast extract, and 2% liquid paraffin. The flasks were then incubated in a shaker sealed with rubber stoppers at 60 °C and 200 rpm. After the liquid paraffin was well emulsified, serial dilutions of the enrichment cultures were plated on a Luria–Bertani (LB) solid medium and incubated at 60 °C. The microbial colonies were chosen and subsequently transferred to fresh liquid paraffin medium. The strains with a good emulsifying activity were selected and stored at −80 °C in LB mixed with sterile glycerol at a final concentration of 20%.

The isolates were classified according to the classification of prokaryote multiphase. Chromosomal DNA was extracted using a MiniBEST Bacterial Genomic DNA Extraction Kit Ver. 2.0 (Takara Biotechnology (Dalian), China) according to the instructions provided by the manufacturer. The universal primers 27f (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1492r (5′-GGTTACCTTGTTACGACTT-3′) were used to amplify microbial 16S rRNA gene fragments.²⁵ The obtained sequences were aligned with the related species on the National Center for Biotechnology Information (NCBI) database, and used to construct phylogenetic trees based on the neighbor-joining method with 1000 bootstrap replicates using MEGA 6.0.²⁶ Subsequently, the phenotypic and physicochemical characteristics of the isolate were identified.

Screening of different types of carbon sources for bioemulsifier production

Considering the efficiency and economic benefit of bioemulsifier production, the potential of the strain A-2 using various carbon sources, particularly inexpensive carbon sources, to produce bioemulsifiers was evaluated. Five types of carbon sources were selected, including water-soluble substrates: glucose, ethanol, and sodium acetate, as well as water-insoluble substrates: hexadecane and olive oil. The strain was incubated in 250 mL flask filled with 100 mL of the fermentation medium containing 4.8 g L⁻¹ K₂HPO₄·3H₂O, 1.5 g L⁻¹ KH₂PO₄, 1 g L⁻¹ (NH₄)₂SO₄, 0.5 g L⁻¹ trisodium citrate, 0.2 g L⁻¹ MgSO₄·7H₂O, 0.1 g L⁻¹ yeast extract, and 1 mL of a trace element solution containing 2 g L⁻¹ CaCl₂·2H₂O, 0.4 g L⁻¹ MnCl₂·4H₂O, 0.4 g L⁻¹ NiCl₂·6H₂O, 0.4 g L⁻¹ ZnSO₄·7H₂O, 0.2 g L⁻¹ FeCl₃·6H₂O, and 0.2 g L⁻¹ NaMoO₄·2H₂O. Moreover, 0.5% (g V⁻¹) of a carbon source was added, and the pH of the medium was adjusted to 7.0 before sterilization. The amount of the inoculation of the fermentation broth was 3%, which was a seed broth grown in LB broth for 7 h at 60 °C. The samples of the culture medium were analyzed periodically for the bacteria concentration, emulsifying activity, amount of residual acetate, and surface tension.

Extraction and purification of bioemulsifier

To remove the cells completely, the culture broth was centrifuged twice at 12 000 rpm, 4 °C, and 30 min. The supernatant was diluted with three volumes of ice-cold acetone for the extraction of bioemulsifier.²⁷ The mixture was stored at 4 °C for 24 h and centrifuged at 9000 rpm, 4 °C for 15 min. The bioemulsifier was obtained as a precipitate, which was collected, lyophilized, and weighed. To purify the bioemulsifier and reduce the content of protein impurities, a three-volume of a mixture of chloroform/n-butanol (4 : 1 v/v) was added to the crude emulsifier solution, mixed vigorously for 5 min, and centrifuged at 9000 rpm for 10 min. The aqueous phase was collected. The above steps were repeated three times. The organic phase was washed with deionized water, centrifuged, and all the aqueous phases were combined. Finally, the solution was dialyzed against distilled water at 4 °C for 72 h and lyophilized. Lyophilized bioemulsifier was subjected to a Sephacryl S-400 column (50 × 2.6 cm, i.d.) and eluted with 0.15 M NaCl at a flow rate of 21 mL h⁻¹. Fractions were collected according to the signal of ultraviolet detector. The bioemulsifier obtained was concentrated, dialyzed against distilled water, and then lyophilized.

Characterization of purified bioemulsifier

Molecular weight determination. The homogeneity and molecular weight of purified bioemulsifier was determined on a Shimadzu LC-20A HPLC system (Kyloto, Japan) equipped with a PL Aquagel-OH MIXED 8 μm column and RID-10 differential refractive index detector. 20 μL of 0.5% bioemulsifier was injected with ultrapure H₂O as the mobile phase at 0.8 mL min⁻¹. The molecular weight was determined by a gel chromatography technique, using dextrans as the standard to generate a standard curve.

Chemical composition. The major components in the bioemulsifier were identified qualitatively by Thin Layer Chromatography (TLC).²⁸ The total carbohydrate content of the purified bioemulsifier was determined by the phenol–sulfuric acid method,²⁹ using glucose as the standard to generate a standard curve. The total protein content was measured by the Kaumas Coomassie brilliant blue staining method using crystalline bovine serum albumin (BSA) as the standard.³⁰ The lipid content of the bioemulsifier was determined by gravimetric estimation.³¹

HPLC analysis (Agilent 1260, USA) was carried out to determine the sugar components and proportions of the purified bioemulsifier. With reference to the literature methods,³² the bioemulsifier was subjected to acid hydrolysis, derivatized with a PMP methanol solution, and finally a 10 μL sample was used for HPLC analysis. The chromatographic conditions were as follows: mobile phase: solvent A: 15% (v/v) acetonitrile + 0.05 M phosphate buffer (KH₂PO₄–NaOH, pH 6.9). Solvent B: 40% (v/v) acetonitrile + 0.05 M phosphate buffer (KH₂PO₄–NaOH, pH 6.9). The gradient mode: time gradient: 0–10–30 min, and the corresponding concentration gradient of solvent B was 0–8–20%. The column temperature was room temperature, the flow rate was 1.0 mL min⁻¹, and the detection wavelength was 250 nm.

The amino acid composition of the sample was measured using a Hitachi L-8800 automatic amino acid analyzer. The polypeptide content of the sample was calculated as follows: first, 1 g of the purified bioemulsifier was subjected to acid hydrolysis for 22 h and filtered. Then, 1 mL sample was evaporated under vacuum, and a volume of 0.02 M HCl was added. The resulting solution was diluted with ultrapure water to 25 mL, left to stand at room temperature for 30 min, and tested.

Functional groups analysis by Fourier transform infrared spectroscopy. For infrared spectrum analysis, 5 mg of the purified bioemulsifier was ground with potassium bromide, compressed into a thin pellet, and analyzed using a FT-IR spectrometer (Bio-rad, USA) in the range 400–4000 cm⁻¹. The functional groups in the bioemulsifier were deduced from the IR spectra.

Determination of emulsifying activity

The emulsifying activity of the cell-free supernatant or bioemulsifier solution was determined as follows: the cell-free supernatant or emulsifier solution was added to a test tube containing equal volumes of hydrocarbons (n-hexane, n-hexadecane, liquid paraffin, xylene, diesel, and crude oil), mixed vigorously for 2 min, and settled at room temperature for 24 h. The emulsification index (E24) was calculated as the ratio of the height of the emulsion layer to the total height of the mixture.³³

Emulsification stability assays

To explore the application range of the bioemulsifier, the effects of environment factors on the stability of the bioemulsifier were evaluated. An experiment was carried out to investigate the effect of pH on the emulsifying activity of the bioemulsifier. The pH of the bioemulsifier solution was adjusted from 1 to 14 using 1.0 M NaOH or 1.0 M HCl. The emulsifying activity of the bioemulsifier was then measured. The salt sensitivity of the bioemulsifier was tested at 5%, 10%, 20%, and 30% salt concentrations. Moreover, we evaluated the emulsification stability at different temperatures (4 °C, 30 °C, 45 °C 60 °C, 70 °C, 80 °C, and 100 °C). Finally, the change in the diameter of oil droplets in the emulsion layer was measured by optical microscopy using a microscopic image analysis software at a certain time interval.³⁴ Diesel was used as the test hydrocarbon.

Core-flooding test

The potential application of the bioemulsifier-producing strain A-2 in MEOR was assessed using the sand-pack core-flooding technique. The experiment simulated a high-temperature reservoir condition. The test temperature was 69 °C, and the pressure was 10 MPa. The test core models were as follows: 600 mm in length, 38 mm in diameter, a porosity of 33.1–33.8%, a permeability of 1.62–1.65 μm², and a crude oil saturation of 92.2% and 93.3% (the ratio of the original oil in place to the volume of brine required to saturate the column was the initial oil saturation, Table 1). The process was as follows: in the first water flooding, the well core was injected with water until the water content of the driven-out liquid was >98%. The oil was displaced continuously until almost no oil was present in the driven-out liquid, which means that the core reached its residual oil saturation. Subsequently, a 0.5 pore volume of the fermentation broth was injected into the water-flooded core, followed by a 15 day shut-in period. Finally, water flooding was performed again until no further oil was observed. The ratio of the volume of crude oil extracted to the volume of the original oil in place could be the percentage of oil recovery. The control was performed under the same conditions but only using water.³⁵

Table 1 The parameters and results of the core-flooding test

Tested project	Pore volume (mL)	Oil saturation (mL)	Water permeability (μm^2)	Oil displacement efficiency (%)
				First water flooding	Successive water flooding	Oil displacement efficiency
Control	230	212.1	1.62	68.4	68.7	0.3%
Strain A-2	225	209.9	1.65	65.7	72.5	6.8%

---

Results and discussion

Identification of enriched facultative anaerobic surfactant-producers

After enrichment, a total of 5 thermophilic strains were isolated. Among them, strain A-2 showed excellent emulsification to liquid paraffin. The strain A-2 was identified by morphological, biochemical, and molecular classification methods. The 16S rDNA sequences (1362 bp) of strain A-2 showed 99% sequence similarity to G. stearothermophilus (Fig. S1; ESI†). The sequence was submitted to GenBank, and its accession number is KR011929. Strain A-2 was a Gram-positive, facultative anaerobic [Fig. S2; ESI†], rod-shaped bacterium with flagella around; the strain could exercise and showed rounded colonies when inoculated on Luria–Bertani (LB) plates. A comparison of the similarities of physiological and biochemical characteristics between the strain A-2 and the species in Geobacillus genus proved that strain A-2 was G. stearothermophilus [Table S1; ESI†].

Effect of carbon sources on bioemulsifier production

The emulsifier production of strain A-2, when growing on fermentation medium containing different carbon sources, was investigated. Fig. 1 showed that all the water-soluble and non-water soluble substrates could support the growth of strain A-2, affording the bioemulsifier production. The bioemulsifier production yields were 3.78 g L⁻¹, 3.03 g L⁻¹, 2.95 g L⁻¹, 2.93 g L⁻¹, and 2.83 g L⁻¹ when sodium acetate, ethanol, glucose, olive oil, and hexadecane were used as the carbon sources, respectively. Notably, the amount of bioemulsifier was higher when 0.5% (g V⁻¹) sodium acetate was used as the carbon source. Although, A-2 was identified as an emulsifier producer, the G. stearothermophilus UCP986 strain was previously reported to be able to produce surfactants, and the yield was 2.3 g L⁻¹ when adding 7.5% palm oil as the carbon source.²⁴ The metabolism characters of strain A-2 using hydrocarbons and organic acids, especially acetate, to produce emulsifier indicate that it is a good candidate for the in situ biosurfactant-mediated MEOR process since the acetate ion is one of the abundant sources of carbon except hydrocarbons in oil reservoirs, which is essential to the survival of microorganisms;³⁶ few bacteria have been reported to use acetate as a carbon source to produce biosurfactants.^37–39

Fig. 1 Effect of the carbon sources on the yield, emulsifying activity of G. stearothermophilus A-2.

To better understand the production of the emulsifier, the growth of strain A-2 and the changes of emulsifying activity were carried out in sodium acetate medium. Fig. 2 shows the cell growth, emulsifying activity, and surface tension of the fermentation broth, along with the substrate consumption. The bacteria reached the maximum bacterial concentration in 6 h, along with the rapid depletion of the substrate. Subsequently, the cells entered the stationary phase, consuming the substrates slowly. The emulsifying activity clearly increased during the exponential phase and reached the maximum activity of ∼61.9% after cultivating for 8 h. There was a positive correlation between the emulsifying activity and bacteria growth. The same feature was observed in several other bioemulsifier-producing microorganisms.^15,40,41 However, the surface tension of the fermentation broth did not decrease significantly, i.e., 63.9 mN m⁻¹ to 54.0 mN m⁻¹. This indicates that the product may be a bioemulsifier, which was more effective in stabilizing oil-in-water emulsions without significantly decreasing the surface tension.

Fig. 2 OD₆₀₀, surface tension, emulsifying activity, and residual acetate profiles of the strain A-2.

Characterization of bioemulsifier

Molecular weight. Only a sharp peak was detected and collected from Sephacryl S-400 column. Then, the purified sample was analyzed with gel-permeation chromatography. The homogeneity of the bioemulsifier was proven by the single and symmetrical peak (Fig. S3; ESI†). The molecular weight of the bioemulsifier was calculated to be approximately 9 × 10⁶ Da, according to the calibration curve with standard dextrans.

Chemical composition. The results of qualitative analysis of sugar, protein and lipid by TLC showed that there were carbohydrate and protein components in the bioemulsifier, but no lipid component (Fig. S4; ESI†). The chemical composition analysis showed that the bioemulsifier produced by G. stearothermophilus A-2 is a glycoprotein, composed of 71.4% sugar and 27.8% protein. The composition of the polysaccharide fraction was analyzed further by high-performance liquid chromatography (HPLC). According to the retention times of the standard samples, the monosaccharides in the bioemulsifier produced by strain A-2 were mannose (33.5%), glucose (30.9%), galactose (29.7%), and glucuronic acid (5.9%) (Fig. 3). Galactose showed two peaks, probably because of tautomerism in the structure of the sample; such tautomers could not be separated and existed in a dynamic equilibrium.

Fig. 3 HPLC analyses of standard monosaccharides and the monosaccharide constituents of bioemulsifier. (A) Mixture of PMP-labeled: (1) mannose; (2) rhamnose; (3) glucose; (4) galactose; (5) glucuronic acid. (B) Sample: (1) mannose; (2) glucose; (3) galactose; (4) glucuronic acid.

Table S2 (ESI†) shows that A-2 bioemulsifier contained 17 types of amino acids. Among these amino acids, the total content of the polar amino acids was 13.74 g/100 g, and that of the nonpolar amino acids was 14.01 g/100 g. The best-studied bioemulsifiers is the emulsan from Acinetobacter RAG-1, which is a complex of an anionic heteropolysaccharide and protein, whose surface activity results from the fatty acids that are attached to the polysaccharide backbone via O-ester and N-acyl linkages. The polysaccharide backbone consists of three amino sugars (D-galactosamine, D-galactosaminouronic acid and a dideoxydiaminohexose); the fatty acid side chains range in length from 10 to 20 carbons, and represent 5–23% (w/w) of the polymer.⁴² Acinetobacter calcoaceticus BD4 produces surface-active extracellular polysaccharide–protein complexes, which consist of L-rhamnose, D-glucose, D-glucuronic and D-mannose.⁴³ Alasan, which is produced by Acinetobacter radioresistens, is a complex of an anionic polysaccharide and protein with a molecular weight of approximately 10⁶ Da,⁴⁴ whereas the polysaccharide component of alasan is unknown. The chemical composition of bioemulsifier produced by strain A-2 was different from the above reported emulsifiers, thus it was a novel glycoprotein emulsifier.

Functional groups and chemical bonds of the bioemulsifier. Fourier transform infrared spectroscopy showed the stretching and deformation vibration of a compound. A-2 bioemulsifier had several characteristic absorption peaks in the range 400–4000 cm⁻¹ (Fig. S5; ESI†). The peak at 1108 cm⁻¹ can be attributed to C–O stretching vibration. The peak at 1400 cm⁻¹ can be assigned to C–H stretching vibration in sugar, and the peak at 1654 cm⁻¹ can be attributed to amide carbonyl stretching vibration. A peak at 3395 cm⁻¹ can be attributed to O–H stretching vibration. The FT-IR spectrum of A-2 bioemulsifier showed the presence of OH and CONH groups, which are characteristic of glycoproteins. The results of the infrared spectrum indicated that the bioemulsifier was a glycoprotein, which was consistent with the results of chemical composition analysis. To the best of our knowledge, strain A-2 is the first strain of G. stearothermophilus that produces a glycoprotein bioemulsifier.

Emulsifying activity

The emulsifying activity is an important characteristic of a bioemulsifier. The emulsifying activity of bioemulsifier produced by strain A-2 for various hydrocarbons was evaluated (Fig. 4). The bioemulsifier produced by strain A-2 with a concentration of 0.5% (g V⁻¹) successfully emulsified pure alkanes and aromatics (n-hexane, hexadecane, and xylene), as well as mixed alkanes and aromatics, such as diesel, olive oil, and crude oil, thereby forming oil-in-water emulsions (i.e. O/W emulsions), wherein the emulsions could reduce significantly the apparent viscosity of in-place oils and improve their mobility in porous media.^45,46 However, some other bioemulsifiers showed considerable substrate specificity.⁴⁷ For example, emulsan, an extensively studied bioemulsifier, did not emulsify pure aliphatic, cyclic or aromatic hydrocarbons, but showed good emulsifying activity for the mixtures of those compounds.^40,48,49 This feature makes the bioemulsifier produced by bacteria A-2 more suitable for wide industrial applications, such as oil-tank cleaning, bioremediation, and oil transportation.

Fig. 4 Emulsification of various hydrocarbon substrates by the bioemulsifier produced by G. stearothermophilus A-2.

Stability of bioemulsifier

The stability of the bioemulsifier was investigated at different temperatures, sodium chloride concentrations, pH, and placing time. Fig. 5A shows that the temperature in the range 4–60 °C did not evidently affect the emulsifying activity. With an increase in temperature, the emulsifying activity decreased slightly; for example, at 80–100 °C, the emulsifying activity was still 57.2–62.3%. This feature makes the bioemulsifier a very good candidate for oil exploration in high-temperature oil reservoirs.

Fig. 5 Effects of temperature, salinity, pH, and placing time on the emulsifying activity of bioemulsifier. (A) Temperature; (B) salinity; (C) pH; (D) placing time.

Fig. 5B shows that the emulsifying activity of the bioemulsifier was still high when the salt concentration was in the range 5–30%. Nevertheless, the chemical surfactants, such as SDS (sodium dodecyl sulfate), Triton X-100, or Tween 80, did not show emulsifying activity at a 10–12% NaCl concentration.^50,51 This confirms that the bioemulsifier produced by G. stearothermophilus A-2 was halotolerant.

The highest emulsifying activity was observed at pH 7–14, whereas it decreased slightly at pH 1–6 (Fig. 5C). This is consistent with the emulsifying activity of the biosurfactant produced by G. stearothermophilus strains X2 and X3, whose emulsifying activity increased at pH 7–12.²³ The bioemulsifier is stable at alkaline pH and exhibit halotolerance; therefore, the bioemulsifier can be used for the bioremediation of oil spills in a marine environment.

The emulsion layer of diesel formed by the bioemulsifier remained stable for 12 months at room temperature, whereas the emulsion layers formed by other biosurfactants showed demulsification after one or three months.^40,48,52 Furthermore, little change was observed within the micron level in the particle size of oil droplets in the emulsion layer (Fig. 5D and S6; ESI†). At 24 h, 1 month and 2 months, mostly small sized (1 μm) of oil droplets and densely packed particles were observed. At 2.5 months, large sized particles appeared, and the particles distribution was in the range of 0.2 μm to 3.1 μm. These particles began to rearrange with time. The large sized particles increased in number due to the coalescence of small sized particles. The particle size distribution was in the range of 1.4 μm to 7.2 μm at 12 month. The O/W emulsion is a metastable system with water as the continuous phase, which ensures that the droplets of oil pass through the porous media at a low viscosity. The coalescence tendency of oil droplets intensifies as the drop size increases. In order to avoid phase separation of oil and water during transportation, the oil drop size in the emulsion should be stabilized to less than 50 μm, and the life-time of the emulsion should be long enough.^45,46 Thus, this excellent emulsion stability makes the bioemulsifier a powerful tool for enhancing oil recovery and the transportation of heavy oils.

Core-flooding test

The sand-pack technique has been used widely to evaluate the biosurfactant potential to enhance oil recovery. The results of the core-flooding tests indicated that G. stearothermophilus A-2 enhanced the oil recovery by 6.8% under simulated high-temperature reservoir conditions (Table 1). Biosurfactants can improve the mobility of oil in porous media through changing rock wettability and emulsifying oil, thus increasing the efficiency of crude oil recovery. Most previous studies have focused on the use of surface-active biosurfactant and the producer for MEOR.⁵³ A polysaccharide–lipid complex bioemulsifier produced by Alcaligenes faecalis increased residual oil recovery by 10.7% in a sand-pack model.⁵⁴ Mohamad Ali Fulazzaky et al. showed that the simulation of MEOR by three MCF scenarios of injecting nutrients, microbial culture and bioproducts would be expected to improve crude oil production, increasing it to approximately 3.09%, 14.27%, and 8.48% of the recovery factor, respectively,⁵⁵ whereas the oil displacement efficiencies of above reported water-flooding step-1 were in the range of 33.10–44.2%. In contrast, the oil displacement efficiency of first water-flooding in this study was 65.4%, which means that the amount of oil remaining in artificial cores was less, similar to depletion-type reservoir, and it was difficult to enhance oil recovery. Thus, the G. stearothermophilus A-2 strain enhanced the oil recovery by 6.8% is efficient at lower oil saturation conditions.

Conclusions

In this study, G. stearothermophilus A-2, a thermophilic, facultative anaerobic, emulsifier-producing strain was isolated. The strain could use sodium acetate as the carbon source to gain a high yield bioemulsifier. The produced bioemulsifier was a novel glycoprotein emulsifier, and emulsified various hydrocarbons successfully at a wide range of temperatures, salinity, pH, and placing times. Notably, the emulsion layer of diesel remained stable for 12 months at room temperature and with little change in particle size. The fermentation broth of strain A-2 increased the oil recovery efficiency by 6.8% in core flooding experiments. These results indicate that the bioemulsifier-producing G. stearothermophilus A-2 is a good candidate for the MEOR technique.

Acknowledgements

This work was supported by the Natural Science Foundation of China (Grant No. 41373074 and 31500414).

References

1. S. Sun, Y. Luo, S. Cao, W. Li, Z. Zhang, L. Jiang, H. Dong, L. Yu and W. M. Wu, Bioresour. Technol., 2013, 144, 44–49.
2. H. Ghojavand, F. Vahabzadeh and A. K. Shahraki, J. Pet. Sci. Eng., 2012, 81, 24–30.
3. A. Singh, N. Parmar and O. Ward, in Geomicrobiology and Biogeochemistry, ed. N. Parmar and A. Singh, Springer, Berlin, Heidelberg, 2014, vol. 39, pp. 239–245.
4. S. Sun, Z. Zhang, Y. Luo, W. Zhong, M. Xiao, W. Yi, L. Yu and P. Fu, Bioresour. Technol., 2011, 102, 6153–6158.
5. R. Sen, Prog. Energy Combust. Sci., 2008, 34, 714–724.
6. I. Lazar, I. G. Petrisor and T. F. Yen, Pet. Sci. Technol., 2007, 25, 1353–1366.
7. P. Sarafzadeh, A. Niazi, V. Oboodi, M. Ravanbakhsh, A. Z. Hezave, S. S. Ayatollahi and S. Raeissi, J. Pet. Sci. Eng., 2014, 113, 46–53.
8. N. Youssef, M. S. Elshahed and M. J. McInerney, in Adv. Appl. Microbiol., Academic Press, 2009, vol. 66, pp. 141–251.
9. A. Singh, J. D. Van Hamme and O. P. Ward, Biotechnol. Adv., 2007, 25, 99–121.
10. S. Lang, Curr. Opin. Colloid Interface Sci., 2002, 7, 12–20.
11. J. Lin, B. Hao, G. Cao, J. Wang, Y. Feng, X. Tan and W. Wang, J. Pet. Sci. Eng., 2014, 122, 354–359.
12. F. Zhao, J. Zhang, R. Shi, S. Han, F. Ma and Y. Zhang, RSC Adv., 2015, 5, 36044–36050.
13. T. Barth, Appl. Geochem., 1991, 6, 1–15.
14. P. Darvishi, S. Ayatollahi, D. Mowla and A. Niazi, Colloids Surf., B, 2011, 84, 292–300.
15. M. Shavandi, G. Mohebali, A. Haddadi, H. Shakarami and A. Nuhi, Colloids Surf., B, 2011, 82, 477–482.
16. I. M. Banat, A. Franzetti, I. Gandolfi, G. Bestetti, M. G. Martinotti, L. Fracchia, T. J. Smyth and R. Marchant, Appl. Microbiol. Biotechnol., 2010, 87, 427–444.
17. E. Rosenberg and Z. E. Ron, Appl. Microbiol. Biotechnol., 1999, 52, 154–162.
18. H. Amani, M. H. Sarrafzadeh, M. Haghighi and M. R. Mehrnia, J. Pet. Sci. Eng., 2010, 75, 209–214.
19. J. F. B. Pereira, E. J. Gudiña, R. Costa, R. Vitorino, J. A. Teixeira, J. A. P. Coutinho and L. R. Rodrigues, Fuel, 2013, 111, 259–268.
20. E. J. Gudiña, J. F. B. Pereira, R. Costa, J. A. P. Coutinho, J. A. Teixeira and L. R. Rodrigues, J. Hazard. Mater., 2013, 261, 106–113.
21. N. Youssef, D. R. Simpson, M. J. McInerney and K. E. Duncan, Int. Biodeterior. Biodegrad., 2013, 81, 127–132.
22. T. N. Nazina, A. A. Grigoryan, G. A. Osipov, T. P. Tourova, S. S. Belyaev, A. E. Ivanova, A. B. Poltaraus, M. V. Ivanov, E. V. Novikova, A. M. Lysenko and V. V. Petrunyaka, Int. J. Syst. Evol. Microbiol., 2001, 51, 433–446.
23. C. Zheng, Z. Li, J. Su, R. Zhang, C. Liu and M. Zhao, J. Appl. Microbiol., 2012, 113, 44–51.
24. A. M. A. T. Jara, R. F. S. Andrade and G. M. Campos-Takaki, Colloids Surf., B, 2013, 101, 315–318.
25. S. S. Gauri, S. M. Mandal, K. C. Mondal, S. Dey and B. R. Pati, Bioresour. Technol., 2009, 100, 4240–4243.
26. K. Tamura, G. Stecher, D. Peterson, A. Filipski and S. Kumar, Mol. Biol. Evol., 2013, 30, 2725–2729.
27. E. J. L. Soon Han Kim, S. Ok Lee, J. Dong Lee and T. H. Lee, Biotechnol. Appl. Biochem., 2000, 31, 249–253.
28. L. Cai, in Current Protocols Essential Laboratory Techniques, John Wiley & Sons, Inc., 2008.
29. M. Dubois, K. A. Gilles, J. K. Hamilton, P. Rebers and F. Smith, Anal. Chem., 1956, 28, 350–356.
30. I. Syrový and Z. Hodný, J. Chromatogr. B: Biomed. Sci. Appl., 1991, 569, 175–196.
31. M. Manocha, G. San-Blas and S. Centeno, J. Gen. Microbiol., 1980, 117, 147–154.
32. S. Honda, E. Akao, S. Suzuki, M. Okuda, K. Kakehi and J. Nakamura, Anal. Biochem., 1989, 180, 351–357.
33. S. M. M. Dastgheib and M. A. Amoozegar, Biotechnol. Lett., 2007, 30, 263–270.
34. P.-K. Gao, G.-Q. Li, L.-X. Zhao, X.-C. Dai, H.-M. Tian, L.-B. Dai, H.-B. Wang, H.-D. Huang, Y.-H. Chen and T. Ma, J. Biosci. Bioeng., 2014, 117, 215–221.
35. P. Gao, G. Li, X. Dai, L. Dai, H. Wang, L. Zhao, Y. Chen and T. Ma, World J. Microbiol. Biotechnol., 2013, 29, 2045–2054.
36. L. Feng, W. Wang, J. Cheng, Y. Ren, G. Zhao, C. Gao, Y. Tang, X. Liu, W. Han, X. Peng, R. Liu and L. Wang, Proc. Natl. Acad. Sci. U. S. A., 2007, 104, 5602–5607.
37. P. Das, S. Mukherjee and R. Sen, Bioresour. Technol., 2009, 100, 1015–1019.
38. R. Makkar and S. S. Cameotra, J. Ind. Microbiol. Biotechnol., 1998, 20, 48–52.
39. R. Makkar and S. S. Cameotra, J. Ind. Microbiol. Biotechnol., 1997, 18, 37–42.
40. E. J. Gudiña, J. F. Pereira, R. Costa, D. V. Evtuguin, J. A. Coutinho, J. A. Teixeira and L. R. Rodrigues, Microb. Cell Fact., 2015, 14, 1–11.
41. A. Franzetti, G. Bestetti, P. Caredda, P. La Colla and E. Tamburini, FEMS Microbiol. Ecol., 2008, 63, 238–248.
42. A. Zuckerberg, A. Diver, Z. Peeri, D. L. Gutnick and E. Rosenberg, Appl. Environ. Microbiol., 1979, 37, 414–420.
43. N. Kaplan, E. Rosenberg, B. Jann and K. Jann, Eur. J. Biochem., 1985, 152, 453–458.
44. S. Navon-Venezia, Z. Zosim, A. Gottlieb, R. Legmann, S. Carmeli, E. Z. Ron and E. Rosenberg, Appl. Environ. Microbiol., 1995, 61, 3240–3244.
45. S. Fournanty, Y. L. Guer, K. E. Omari and J. P. Dejean, J. Dispersion Sci. Technol., 2008, 29, 1355–1366.
46. D. Langevin, S. Poteau, I. Hénaut and J. F. Argillier, Oil Gas Sci. Technol., 2004, 59, 511–521.
47. E. Z. Ron and E. Rosenberg, Curr. Opin. Biotechnol., 2002, 13, 249–252.
48. M. A. Luna-Velasco, F. Esparza-García, R. O. Cañízares-Villanueva and R. Rodríguez-Vázquez, Process Biochem., 2007, 42, 310–314.
49. E. Rosenberg, A. Perry, D. T. Gibson and D. L. Gutnick, Appl. Environ. Microbiol., 1979, 37, 409–413.
50. T. de Sousa and S. Bhosle, Bioresour. Technol., 2012, 123, 256–262.
51. A. Monteiro, M. Bonfim, V. Domingues, A. Corrêa, E. Siqueira, C. Zani and V. Santos, Bioresour. Technol., 2010, 101, 5186–5193.
52. S. Dastgheib, M. Amoozegar, E. Elahi, S. Asad and I. Banat, Biotechnol. Lett., 2008, 30, 263–270.
53. R. Marchant and I. M. Banat, Trends Biotechnol., 2012, 30, 558–565.
54. H. Salehizadeh and S. Mohammadizad, Iran. J. Biotechnol., 2009, 7, 216–223.
55. M. Fulazzaky, D. I. Astuti and M. A. Fulazzaky, RSC Adv., 2015, 5, 3908–3916.

---

Footnotes

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra15626f

‡ Equal contributors.

---