Biotechnology and Bioprocess Engineering 2009, 14: 811-818 DOI/10.1007/s12257-008-0306-y
=
Direct Purification of Burkholderia Pseudomallei Lipase from Fermentation Broth Using Aqueous Two-Phase Systems Chien Wei Ooi1, Beng Ti Tey2, Siew Ling Hii3, Arbakariya Ariff4, Ho Shing Wu5, John Chi Wei Lan5, Ruey Shin Juang5, Siti Mazlina Mustapa Kamal1, and Tau Chuan Ling1* 1
Department of Process and Food Engineering, Universiti Putra Malaysia, 43400 UPM Serdang, Selangor, Malaysia 2 Department of Chemical and Environmental Engineering, Universiti Putra Malaysia, 43400 UPM Serdang, Selangor, Malaysia 3 Faculty of Engineering and Science, Universiti Tunku Abdul Rahman, 53300 Setapak, Kuala Lumpur, Malaysia 4 Faculty of Biotechnology and Biomolecular Sciences, Universiti Putra Malaysia, 43400 UPM Serdang, Selangor, Malaysia 5 Department of Chemical Engineering and Materials Science, College of Engineering, Yuan Ze University, Taiwan Abstract An aqueous two-phase purification process was employed for the recovery of _ìêâÜçäÇÉêá~=éëÉìÇçã~ääÉá=lipase from fermentation broth. The partition behavior of _K=éëÉìÇçã~ääÉá=lipase was investigated with various parameters such as phase composition, tie-line length (TLL), volume ratio (VR), sample loading, system pH, and addition of neutral salts. Optimum conditions for the purification of lipase were obtained in polyethylene glycol (PEG) 6000-potassium phosphate system using TLL of 42.2% (w/w), with VR of 2.70, and 1% (w/w) NaCl addition at pH 7 for 20% (w/w) crude load. Based on this system, the purification factor of lipase was enhanced to 12.42 fold, with a high yield of 93%. Hence, the simplicity and effectiveness of aqueous two-phase systems (ATPS) in the purification of lipase were proven in this study. © KSBB = = = =
hÉóïçêÇëW=~èìÉçìë=íïçJéÜ~ëÉ=ëóëíÉãëI=äáé~ëÉI=éìêáÑáÅ~íáçåI=éçäóÉíÜóäÉåÉ=ÖäóÅçäI=êÉÅçîÉêó=
INTRODUCTION Lipases or triacylglycerol hydrolases (E.C. 3.1.1.3) are hydrolases that catalyze the hydrolysis of carboxyl ester bonds present in triacylglycerols [1]. B. pseudomallei lipase was ascribed in the lipolytic enzyme family that encodes an operon together with Lifs (lipase specific foldases) [2]. This feature enables the folding of lipases into an enzymatically active form and the secretion of these active lipases into the surrounding medium [3,4]. B. pseudomallei lipase is well known for its characterizations such as high temperature stability, organic solvent tolerance, and high catalytic activity for various substrates [2]. The wide-ranging versatility of lipases has become a great interest to industrial applications [5] such as food technology, detergent, bioenergy [6], pharmaceutical industry [7,8], and biodiesel fuel [9]. Microbial lipases have been purified by conventional pro*Corresponding author Tel: +60-3-89466360 Fax: +60-3-86567123 e-mail:
[email protected]
cedures including precipitation or chromatography [10], with partial purification and low yields. These enzyme purification processes present a significant problem that mainly derived from the complexity of the protein mixtures and the necessity to retain their biological activity. ATPS is formed by dissolving the water-soluble phase components beyond a critical concentration, in which two immiscible phases are formed [11]. ATPS generates a specific environment that selectively partition and concentrate target biomolecules into one of the phases, while maintaining the native structure of biomolecules since both phases of ATPS are predominantly water-based (80~85%). Extraction using ATPS offers a better alternative to replace numerous steps in conventional downstream processing, such as extraction, clarification, concentration, and intermediate purification [12-14]. Moreover, equilibrium mass transfer in ATPS is easily reached and separation could be selective and rapid [15]. Apart from that, ATPS purification is suitable to be incorporated into large-scale application prior to the ease of process scaling-up and the capability of continuous operation [16,17]. In present, there is no available information regarding the
812
ATPS of this extracellular B. pseudomallei lipase. Therefore, a detail study was made with an aim of applying ATPS extraction for B. pseudomallei fermentation culture. The polymer-salt system (PEG-phosphate) was selected owing to the inexpensive phase forming chemicals and the lower phase viscosity, as compared to the other polymer-polymer system, such as PEG-dextran system. The dependence of lipase partition behavior on variables such as molecular weight of polymer, TLL, VR, pH, and addition of salts was studied in this research. These parameters could be manipulated in order to achieve the most favorable condition of ATPS extraction.
MATERIALS AND METHODS Materials
PEG 3000, 6000, and 8000 g/mol and p-nitrophenyl laurate (pNPL) were obtained from Fluka. Co. (USA). Dipotassium hydrogen phosphate (K2HPO4) and potassium dihydrogen phosphate (KH2PO4) were sourced from Merck (Darmstadt, Germany). Bicinchoninic acid (BCA) solution was purchased from Sigma Aldrich (St. Louis, USA). All chemicals were of analytical grade. Feedstock Preparation
B. pseudomallei was grown in batch culture medium containing the constituents: 0.33% (w/v), nutrient broth; 0.10% (w/v), CaCl2; 1.00% (v/v), Tween 80; and 1.00% (w/v), gum arabic. The pH value of the medium was adjusted to 9.0. After 5% (v/v) inoculation, the culture was incubated at 37oC and agitated at 250 rpm. After 72 h of incubation, the culture was harvested without clarification and was directly used for the ATPS experiments. Enzyme Activity Assay
The assay was performed as described previously [18,19]. The reaction mixture contained 25 µL, enzyme extract; 200 µL, 0.05 M phosphate buffer (pH 6.5); 5 µL, Triton-X; and 25 µL, 0.02 M pNPL in ethanol. The amount of p-nitrophenol released in the hydrolysis of pNPL was measured at absorbance of 405 nm using a microplate reader (Tecan Sunrise). One unit of enzymatic activity was defined as the amount of lipase enzyme needed to release 1 µmol pNPL per minute. BCA Assay
The total protein concentrations in the selected samples were analyzed using BCA assay [20]. A volume of 50 µL of sample was mixed with 200 µL of the working reagent in microtiter plate. The mixture was incubated at 37oC (30 min). The absorbance at 562 nm was measured against a reagent blank, which contains an appropriate diluted phase solution without protein sample.
Phase Diagrams of Various ATPS
The binodal curves were estimated using cloud point method as described by Albertsson [15]. TLL describes the compositions of the two phases, which are in equilibrium and it was calculated as: TLL = ΔP 2 + ΔC 2
(1)
where ∆P and ∆C are the difference between PEG and phosphate concentration, respectively, in the two phases. The concentration of PEG and salts were analyzed by refractive index and conductivity measurement, respectively [15]. The salt concentration was determined from a calibration curve that had been constructed with a range of standard salt concentrations. PEG concentration in the phase was determined by subtracting the refractive index value, which was contributed by salts. Preparation of ATPS
Concentration of PEG stock solutions was 50% (w/w) polymer by mass. Different stock solutions 40% (w/w) of K2HPO4 and KH2PO4 were used to prepare the ATPS at different pHs. The ATPS were prepared in 15 mL graduated centrifuge tubes by weighting an appropriate amount of PEG stock solution, stock solution of phosphate at desired pH, and crude feedstock at 20% (w/w). To the mixture, adequate distilled water was added to achieve a final mass of 10 g. The mixtures were mixed thoroughly by gentle agitation until equilibrium. The phase separation was accelerated by centrifugation at 4,000 × g for 10 min. The volumes of both top and bottom phases were recorded. The samples were analyzed for lipase activity and total protein concentrations. Partition Coefficient (K), Specific Activity (SA), VR, Purification Factor (PFT), and Yield
The K of the lipase was calculated as the ratio of the lipase activity in the two phases AT (2) AB where AT and AB are the lipase activities in Unit/mL in the top phase and bottom phase, respectively. The SA was defined as the ratio between the enzyme activity (U) in the phase sample and the total protein concentration (mg) K=
SA (U/mg) =
Enzyme activity (U) [Protein] (mg)
(3)
The selectivity (S) was defined as the ratio of the lipase enzyme partition coefficient (Ke) to the protein partition coefficient (Kp)
S=
Ke Kp
(4)
Biotechnol. Bioprocess Eng. 813=
A
B
C
Fig. 1. Phase diagram for PEG and potassium phosphate. The binodal curves for (A) PEG 3000, (B) PEG 6000, and (C) PEG 8000 were plotted against potassium phosphate. Tie-lines in varying length were shown and the composition of ATPS with phase ratio of approximately 1 was indicated as (●) in each of the tie-line.
The VR was defined as the ratio of volume in the top phase (VT) to that in the bottom phase (VB) VR =
VT VB
(5)
The PFT was determined as the ratio of the lipase specific activity in the top phase to the original lipase specific activity in the crude feedstock
PFT =
SA of phase sample SA of crude feedstock
Yield of lipase in top phase was estimated using 100 YT (%) = 1 + (1 / VR × K )
(6)
(7)
Sodium Dodecyl Sulfate-polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis Protein samples were analyzed with SDS-PAGE as de-
scribed previously [21]. Protein samples were concentrated and precipitated using tricholoacetic acid (TCA) solution (10%) before the SDS-PAGE analysis. Acrylamide gel made up of a 12% of resolving gel and a stacking gel of 4.5% was used in this study. The gel was stained with 0.05% (v/v) Coomassie® Brilliant Blue G-250.
RESULTS AND DISCUSSION Effect of PEG Molecular Weight and TLL on Lipase Partitioning
The phase diagrams for PEG-phosphate system with comparable TLLs were shown in Figs. 1A, 1B, and 1C. These systems were built using three different molecular mass of PEG (i.e. 3000, 6000, and 8000), with an increasing trend of the TLL while keeping VR = 1.0 and pH = 7.0 at constant. The logarithmic partition coefficient log K for lipase was shown in Table 1. Majority of the lipases showed a topphase preference, with the partition coefficient of lipase slightly increased at the elevating TLL. Such phenomenon
814
Table 1. Influence of the PEG molecular weight and TLL=on the partitioning of lipase PEG molecular weight 3000
6000
8000
TLL
Log K
Selectivity
30.7
0.26
1.72
34.9
0.53
2.21
37.7
0.43
2.03
40.7
1.28
12.35
43.4
1.13
9.00
46.0
1.98
58.21
48.4
0.96
5.29
50.7
0.90
4.40
29.6
0.58
4.03
32.6
0.60
4.10
35.2
0.92
7.42
37.9
0.91
7.64
40.0
2.26
166.23
42.2
2.48
243.94
44.2
1.18
16.44
46.6
0.88
6.77
27.3
0.00
1.94
30.9
0.47
4.37
33.5
0.33
3.70
36.1
0.73
8.40
38.4
1.25
20.36
40.7
0.74
6.06
42.6
0.63
4.04
44.4
0.55
3.64
The influences of PEG molecular weight and concentration on lipase partitioning were investigated. The log K of PEG 3000, PEG 6000, and PEG 8000 were determined by Equation 2. The results were expressed as a mean of triplicate readings with an estimated error of ± 10%.
had been reported by the other authors [22]. In general, a similar trend of partition efficiency was exhibited for ATPS with different molecular weight of PEG, in which the optimum partitioning of lipase was noticed at long TLL of 40~48% (w/w). In the case of PEG 6000/phosphate ATPS, long TLL at 42.2% (w/w) resulted a significant maximum lipase partitioning at the top phase, with the log K of 2.48. The other systems comprised of PEG 3000 and PEG 8000 had achieved their maximum lipase partitioning at log K of 1.98 (TLL 46.0% (w/w)) and 1.25 (TLL 38.4% (w/w)), respectively. Furthermore, the selectivity of PEG 6000 ATPS is the highest (244), as compared to PEG 3000 (58.21) and PEG 8000 (20.36). It appeared that the low partition efficiency of ATPS with PEG 8000 could be caused by the effect of excluded volume [23]. Precipitation at the interphase was observed since the protein saturation point in the polymer phase had reached. Moreover, an increase in the chain length of PEG will also cause reduction of available free volume (excluded volume) to accommodate lipase in the upper phase [24,25]. Another
tendency that affects the partitioning, is the high PEG molecular mass which will strengthen the viscosity of system and thus unsuitable for processing. A maximum partitioning of lipase at 42.2% (w/w) TLL for PEG 6000 was observed, revealing that a fine balance of both PEG hydrophobicity [26] and salting-out ability of salts was greatly achieved. This suggested that the protein partitioning at higher TLL could lead to extreme Kvalues and purification [27]. As further increase in the molecular weight of PEG, hydroxyl groups for the polymer tend to be lesser and eventually the hydrophobicity of the polymer-rich top phase will be higher [28]. Besides, an increase in the TLL corresponds to higher concentration of salt in the bottom phase. This higher salt concentration will have an impact on the hydrophobic interactions between the proteins and polymer phase, since salt ions will interact with the oppositely charged groups in the protein and dehydrate the proteins by forming a double layer of ionic groups. This will dehydrate the proteins as the hydration effect of salt molecules caused the hydrophobic zones of the protein to be exposed [29]. Hence, the hydrophobic interaction between protein and the PEG molecules was improved in this way [30]. Moreover, the enzyme partition behavior was also associated with the lower free volume available in the bottom phase as the TLL increases, therefore driving most of the lipases toward the polymer-rich top phase [24,31]. As a result, PEG 6000phosphate system with TLL of 42.2% (w/w) was selected to further study the effect of other parameters in ATPS for partitioning lipase. Effect of VR on Lipase Partitioning
The differential partitioning of lipase for TLL 42.2% (w/w) at different phase volume ratios was shown in Fig. 2. Lipase partition coefficients were found increasing gradually in accordance with the increasing VR. The similar trend of enzyme protein partitioning behavior was observed in other findings [32]. In principle, the protein partition behavior is not affected by altering the VR, i.e. by moving along a tieline [33,34] because the relative partitioning of the individual protein will not change. The fact that VR affected the purification was noted in view of the presence of protein precipitation at the interface, as the top-phase volume decreased. The free volume in the top phase was greatly reduced for ATPS with lower VR, leading to the losses of lipases that remained at the top phase. The differences in log K value for each of the ATPS VR are insignificant. Thus, it was very difficult to evaluate the impact of parameters upon the partition behavior of lipase by monitoring partition coefficient only. As a result, the PFT (top phase) and selectivity of both the system were also evaluated. It was found that system with VR of 2.70 showed the best partition efficiency as compared to the other systems. The selectivity and purification fold of VR = 2.70 were 18.01 and 5.86, respectively (Fig. 2). The results indicated that the ATPS with VR at 2.70 gave a higher purification of lipase and therefore optimum for lipase partitioning.
Biotechnol. Bioprocess Eng. 815=
Table 2. Influence of the crude feedstock load on the partitioning of lipase Crude load (% w/w)
Selectivity
Yield (%)
10
2.26
90.44
20
5.56
92.56
30
3.41
81.60
40
4.66
78.53
The table summarizes the extraction results achieved in phase system composed PEG 6000 with TLL 42.2% (w/w). ATPS were comprised of crude feedstock load ranging from 10 to 40% (w/w). The results were expressed as a mean of triplicate readings with an estimated error of ± 10%.
Fig. 2. Influence of the VR on the partitioning of lipase. The partition behavior of lipase with increasing volume ratios was investigated. By selecting several points along the TLL of 42.2% (w/w) gradually, the VR could be varied from 0.31 to 4.00. (♦), the logarithm partition coefficient log K; (▲), selectivity; and (■), PFT of lipase at top phase were calculated and plotted against the VR. The results were expressed as a mean of triplicate readings with an estimated error of ± 10%.
Effect of Crude Load on Lipase Partitioning
An increment in crude feedstock load would be advantageous in the recovery process of larger feedstock volumes by the ATPS. The impact of the loaded mass on the partition of enzymes is important, as the loaded feed stock can alter the phase VR [35] and the partition behavior of target protein [36]. The increasing amounts of both lipases and contaminants in the systems could result in decrease of the ATPS performance. ATPS experiments were carried out by varying the crude load up to 40% (w/w). Table 2 illustrates the effect of crude load on top lipase recovery. Based on the results, crude feedstock load of 20% (w/w) is the maximum capacity on the basis of 10 g ATPS. The selectivity and yield for the 20% (w/w) crude load ATPS was 5.56 and 92.6%, respectively. Higher amount of the sample loading into the ATPS will decrease the volume ratio and affect the composition of ATPS. It appeared that the components in the crude stock had changed the properties of the ATPS; hence the ATPS was not optimum for purification of lipase. Such behavior can be explained by the increasing accumulation of precipitate at the interface, showing that the loss of lipases together with other contaminants in the purification. Therefore it is clear that a 20% of sample loading would be feasible for the maximum top phase recovery of lipase from the crude extract. Effect of pH on Lipase Partitioning
The influence of pH on selectivity and yield of lipase in phase system was shown in Fig. 3. Based on the result, an average of 92% yield was shown in a pH range of 6.5~8.5. The yield and lipase activity was severely reduced when assayed at pH less than 6.5, with most of the lipases partitioned to the bottom phase. This dramatic change of the partition behavior of lipase was caused by the influence of the
Fig. 3. Influence of the pH on the partitioning of lipase. In all experiments, the pH of ATPS was varied between 5.0 and 8.5. The selectivity and yield were calculated using equations 4 and 7, accordingly. The results were expressed as a mean of triplicate readings with an estimated error of ± 10%.
protein charge. As the pH of the system changes, lipase will be partitioned according to the net charge of protein and surface properties other than the charge. Lipase has a pH about 6.3 [19], so at pH 6.5 the lipase is slightly negatively charged and the partitioning will mostly depend on the surface properties rather than the net charge. The lipase is negatively charged at above pH 7~8. PEG tends to act as positively charged [26,37] and interact with the lipase. This interaction improved the partitioning of lipases, as the highest selectivity was achieved at neutral pH 7 (10.98). Furthermore, the yield at pH 7 is the highest (93%). ATPS of pH 7 was selected for further study. Effect of NaCl on Lipase Partitioning=
Partitioning of lipase as a function of varying NaCl concentration [1~7% (w/v)] was studied for TLL 42.2% (w/w) at pH 7 and the result was depicted in Fig. 4. Variation in the salt type and concentration will cause an electrical potential difference between the two phases, as a result of the preference of ions in the one phase relative to the other [38]. This electrostatic potential difference will strongly affect the partitioning of charged protein. The addition of NaCl or other neutral salts in ATPS can affect the water structure and hy-
816
Fig. 4. Influence of the addition of NaCl on the partitioning of lipase. All the scouting experiments were carried out with PEG 6000/potassium phosphate, TLL 42.2% (w/w) at pH 7, and VR of 2.70. (■), selectivity and (♦), PFT were calculated as a function of the NaCl concentrations. The results were expressed as a mean of triplicate readings with an estimated error of ± 10%.
drophobic interactions differently [15], in which the interaction between hydrophobic chain (ethylene group) of PEG and hydrophobic surface area of the lipase will be facilitated. Based on the results, an increase in the enzyme selectivity was induced by 1~2% (w/w) of NaCl. The highest selectivity (8.12) and purification fold of lipase (12.42) were achieved at the addition of 1% (w/w) NaCl. The addition of NaCl above 2% (w/w) will decrease the partition efficiency, as the chemical potential of the solute is affected due to the unequal partitioning of neutral salts between the two phases [39]. Recovery of Lipase from ATPS Extraction
The optimum recovery of lipase from crude feedstock was achieved in ATPS composed of PEG 6000, 42.2% (w/w) TLL with VR = 2.70, 1% (w/w) NaCl addition and 20% crude feedstock load at pH 7.0. The SDS-PAGE profile [21] of the crude feedstock, standard protein markers and purified lipase samples was shown in Fig. 5. The crude feedstock contained a large number of bands (lanes 3 and 4). The bottom phase samples showed lesser and fainter bands than crude extract (lanes 6 and 7). The sample taken from the top phase with highest lipolytic activity and purity showed two prominent dark bands (lane 5), with molecular masses of 66.5 and 40.3 kDa. B. pseudomallei lipases (class 3) in general have molecular mass of about 39.5 kDa [40,41]. The other band (66.5 kDa) might be dimer or aggregate of the lipase, since these two corresponding protein bands were observed from the SDS PAGE analysis of lipase that purified with sucrose gradient ultracentrifugation and ion exchange chromatography (unpublished data).
CONCLUSION The potential of ATPS for the B. pseudomallei lipase puri-
Fig. 5. SDS-PAGE analyses on the recovery of lipase. The purity of partitioned lipase was assessed by 12% SDSPAGE analysis [21]. Molecular weight of standard protein marker ranged 7~175 kDa. SDS-PAGE: lanes 1 and 2, protein molecular markers; lanes 3 and 4, crude feedstock; lane 5, ATPS top phase; and lanes 6 and 7, ATPS bottom phase.
fication in single-step strategy was proven to be suitable for a commercial recovery process. PEG 6000-potassium phosphate system comprised of 42.2% (w/w) TLL, VR of 2.70, 1% (w/w) of NaCl addition, and 20% crude feedstock load at pH 7.0 has provided the optimum conditions with a yield of 93% (purification fold of 12.42). It was shown that molecular mass of PEG, TLL, and VR influenced the lipase partitioning. The addition of salts and modification of pH further facilitated the partitioning of lipase into the top phase. The findings reported here led to an alternative process option of efficient and inexpensive lipase purification, which can be greatly achieved by using PEG-phosphate ATPS. Acknowledgment This study was supported by EScience Fund (03-01-04-SF0785) from the Ministry of Science, Technology and Innovation (MOSTI) and RUGS 91173 (UPM), Malaysia.
Received December 15, 2008; accepted May 13, 2009
REFERENCES 1. Jaeger, K.-E., S. Ransac, B. W. Dijkstra, C. Colson, M. van Heuvel, and O. Misset (1994) Bacterial lipases. FEMS Microbiol. Rev. 15: 29-63. 2. Yang, J., D. Guo, and Y. Yan (2007) Cloning, expression, and characterization of a novel thermal stable and short-chain alcohol tolerant lipase from Burkholderia cepacia strain G63. J. Mol. Catal. B: Enzym. 45: 91-96.
Biotechnol. Bioprocess Eng. 817=
3. El Khattabi, M., P. Van Gelder, W. Bitter, and J. Tommassen (2000) Role of the lipase-specific foldase of Burkholderia glumae as a steric chaperone. J. Biol. Chem. 275: 26885-26891. 4. Rosenau, F. and K.-E. Jaeger (2000) Bacterial lipases from Pseudomonas: regulation of gene expression and mechanisms of secretion. Biochimie. 82: 1023-1032. 5. Saxena, R. K., P. K. Ghosh, R. Gupta, W. S. Davidson, S. Bradoo, and R. Gulati (1999) Microbial lipases: potential biocatalysts for the future industry. Curr. Sci. 77: 101-115. 6. Noureddini, H., X. Gao, and R. S. Philkana (2005) Immobilized Pseudomonas cepacia lipase for biodiesel fuel production from soybean oil. Bioresour. Technol. 96: 769-777. 7. Jaeger, K.-E., B. W. Dijkstra, and M. T. Reetz (1999) Bacterial biocatalysts: molecular biology, three-dimensional structures, and biotechnological applications of lipases. Annu. Rev. Microbiol. 53: 315-351. 8. Pandey, A., S. Benjamin, C. R. Soccol, P. Nigam, N. Krieger, and V. T. Soccol (1999) The realm of microbial lipases in biotechnology. Biotechnol. Appl. Biochem. 29: 119-131. 9. Shimada, Y., Y. Watanabe, T. Samukawa, A. Sugihara, H. Noda, H. Fukuda, and Y. Tominaga (1999) Conversion of vegetable oil to biodiesel using immobilized Candida antarctica lipase. J. Am. Oil Chem. Soc. 76: 789-793. 10. Gupta, R., N. Gupta, and P. Rathi (2004) Bacterial lipases: an overview of production, purification, and biochemical properties. Appl. Microbiol. Biotechnol. 64: 763-781. 11. Zaslavsky, B. Y. (1995) Aqueous two-phase partitioning: physical chemistry and bioanalytical applications. pp. 221-290. Marcel Dekker Inc., NY, USA. 12. Porto, T. S., G. M. Medeiros e Silva, C. S. Porto, M. T. H. Cavalcanti, B. B. Neto, J. L. Lima-Filho, A. Converti, A. L. F. Porto, and A. Pessoa Jr (2008) Liquid-liquid extraction of proteases from fermented broth by PEG/citrate aqueous two-phase system. Chem. Eng. Processing: Process Intensification 47: 716-721. 13. Rosa, P. A. J., A. M. Azevedo, I. F. Ferreira, J. de Vries, R. Korporaal, H. J. Verhoef, T. J. Visser, and M. R. Aires-Barros (2007) Affinity partitioning of human antibodies in aqueous two-phase systems. J. Chromatogr. A 1162: 103-113. 14. Madhusudhan, M. C., K. S. M. S. Raghavarao, and S. Nene (2008) Integrated process for extraction and purification of alcohol dehydrogenase from Baker’s yeast involving precipitation and aqueous two phase extraction. Biochem. Eng. J. 38: 414-420. 15. Albertsson, P. A. (1986) Partition of Cell Particles and Macromolecules. 3rd ed., pp. 1-3. Wiley Interscience, NY, USA. 16. Costa, M. J. L., M. T. Cunha, J. M. S. Cabral, and M. R. Aires-Barros (2000) Scale-up of recombinant cutinase recovery by whole broth extraction with PEG-phosphate aqueous two-phase. Bioseparation 9: 231-238.
17. Roger, G. H. (1994) Liquid-liquid extraction. In: W. M. Courtney (ed.). Protein Purification Process Engineering. Marcel Dekker Inc., NY, USA. 18. Gupta, N., P. Rathi, and R. Gupta (2002) Simplified para-nitrophenyl palmitate assay for lipases and esterases. Anal. Biochem. 311: 98-99. 19. Ishimoto, R., M. Sugimoto, and F. Kawai (2001) Screening and characterization of trehalose-oleate hydrolyzing lipase. FEMS Microbiol. Lett. 195: 231-235. 20. Smith, P. K., R. I. Krohn, G. T. Hermanson, A. K. Mallia, F. H. Gartner, M. D. Provenzano, E. K. Fujimoto, N. M. Goeke, B. J. Olson, and D. C. Klenk (1985) Measurement of protein using bicinchoninic acid. Anal. Biochem. 150: 76-85. 21. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680-685. 22. Bradoo, S., R. K. Saxena, and R. Gupta (1999) Partitioning and resolution of mixture of two lipases from Bacillus stearothermophilus SB-1 in aqueous two-phase system. Process Biochem. 35: 57-62. 23. Almeida, M. C., A. Venâncio, J. A. Teixeira, and M. R. Aires-Barros (1998) Cutinase purification on poly (ethylene glycol)-hydroxypropyl starch aqueous two-phase systems. J. Chromatogr. B: Biomed. Sci. Appl. 711: 151159. 24. Grossman, P. D. and J. L. Gainer (1988) Correlation of aqueous two-phase partitioning of proteins with changes in free volume. Biotechnol. 4: 6-11. 25. Spelzini, D., B. Farruggia, and G. Picó (2005) Features of the acid protease partition in aqueous two-phase systems of polyethylene glycol-phosphate: chymosin and pepsin. J. Chromatogr. B 821: 60-66. 26. Huddleston, J. G., K. W. Ottomar, D. M. Ngonyani, and A. Lyddiatt (1991) Influence of system and molecular parameters upon fractionation of intracellular proteins from Saccharomyces by aqueous two-phase partition. Enzym. Microb. Technol. 13: 24-32. 27. Johansson, G. (1994) Partitioning procedures and techniques: small molecules and macromolecules. Methods Enzymol. 228: 28-42. 28. Saravanan, S., J. R. Rao, T. Murugesan, B. U. Nair, and T. Ramasami (2007) Partition of tannery wastewater proteins in aqueous two-phase poly (ethylene glycol)magnesium sulfate systems: effects of molecular weights and pH. Chem. Eng. Sci. 62: 969-978. 29. Bonomo, R. C. F., L. A. Minim, J. S. R. Coimbra, R. C. I. Fontan, L. H. Mendes da Silva, and V. P. R. Minim (2006) Hydrophobic interaction adsorption of whey proteins: effect of temperature and salt concentration and thermodynamic analysis. J. Chromatogr. B 844: 6-14. 30. Vojdani, F. (1996) Solubility. pp. 11-60. In: G. M. Hall (ed.). Methods of Testing Protein Functionality. Blackie Academic and Professional, London, UK. 31. Rito-Palomares, M. and M. Hernandez (1998) Influence of system and process parameters on partitioning of cheese whey proteins in aqueous two-phase systems. J. Chromatogr. B: Biomed. Sci. Appl. 711: 81-90.
818
32. Cunha, M. T., M. R. Aires-Barros, and J. M. S. Cabral (2003) Extraction for rapid protein isolation. pp. 321372. In: R. Hatti-Kaul and B. Mattiasson (eds.). Isolation and purification of proteins. Marcel Dekker Inc., NY, USA. 33. Walter, H. and G. Johansson (1994) Aqueous two-phase systems, Methods in Enzymology. 2nd ed. Academic Press, London, UK. 34. Hustedt, H., K. H. Kroner, and M. R. Kula (1985) Applications of phase partitioning in biotechnology. pp. 529-587. In: H. Walter, D. E. Brooks, and D. Fischer (eds.) Partitioning in Aqueous Two Phase Systems. Theory, Methods, Uses, and Applications to Biotechnology. Academic Press, Orlando, FL, USA. 35. Abbott, N. L. and T. A. Hatton (1988) Liquid-liquid extraction for protein separations. Chem. Eng. Prog. 84: 31-41. 36. Walter, H., D. E. Brooks, and D. Fisher (1985) Partitioning in aqueous-two phase systems: theory, methods, uses, and application in biotechnology. Academic Press, Orlando, FL, USA. 37. Tanuja, S., N. D. Srinivas, K. S. M. S. Raghava Rao,
38.
39.
40.
41.
and M. K. Gowthaman (1997) Aqueous two-phase extraction for downstream processing of amyloglucosidase. Process Biochem. 32: 635-641. Johansson, G. (1970) Partition of salts and their effects on partition of proteins in a dextran-poly(ethylene glycol)-water two-phase system. Biochim. Biophys. Acta 221: 387-390. Forciniti, D. (2000) Studying the influence of salts on partitioning of proteins. pp. 23-33. In: R. H. Kaul (ed.). Methods in Biotechnology 11: Aqueous Two phase system: Methods and protocols. Humana Press Inc., Totowa, NJ, USA. Mahfouz, M. E., T. H. Grayson, D. A. B. Dance, and M. L. Gilpin (2006) Characterization of the mrgRS locus of the opportunistic pathogen Burkholderia pseudomallei: temperature regulates the expression of a two-component signal transduction system. BMC Microbiol. 6: 70. Yeo, S. H., T. Nihira, and Y. Yamada (1998) Purification and Characterization of tert-Butyl Ester-hydrolyzing Lipase from Burkholderia sp. YY62. Biosci. Biotechnol. Biochem. 62: 2312-2317.
