Novel lipase purification methods - a review of the latest developments.

Biotechnology Journal

Biotechnol. J. 2015, 10

DOI 10.1002/biot.201400301

www.biotechnology-journal.com

Review

Novel lipase purification methods – a review of the latest developments Chung Hong Tan1, Pau Loke Show1,2, Chien Wei Ooi3, Eng-Poh Ng4, John Chi-Wei Lan5 and Tau Chuan Ling6 1 Department

of Chemical and Environmental Engineering, University of Nottingham Malaysia Campus, Jalan Broga, Selangor Darul Ehsan, Malaysia 2 Manufacturing and Industrial Processes Division, Centre for Food and Bioproduct Processing, University of Nottingham Malaysia Campus, Jalan Broga, Selangor Darul Ehsan, Malaysia 3 Chemical Engineering, Monash University Malaysia, Selangor, Malaysia 4 School of Chemical Sciences, Universiti Sains Malaysia, Minden, Malaysia 5 Department of Chemical Engineering and Material Science, Yuan Ze University, Chungli, Taoyuan, Taiwan 6 Institute of Biological Sciences, University of Malaya, Kuala Lumpur, Malaysia

Microbial lipases are popular biocatalysts due to their ability to catalyse diverse reactions such as hydrolysis, esterification, and acidolysis. Lipases function efficiently on various substrates in aqueous and non-aqueous media. Lipases are chemo-, regio-, and enantio-specific, and are useful in various industries, including those manufacturing food, detergents, and pharmaceuticals. A large number of lipases from fungal and bacterial sources have been isolated and purified to homogeneity. This success is attributed to the development of both conventional and novel purification techniques. This review highlights the use of these techniques in lipase purification, including conventional techniques such as: (i) ammonium sulphate fractionation; (ii) ion-exchange; (iii) gel filtration and affinity chromatography; as well as novel techniques such as (iv) reverse micellar system; (v) membrane processes; (vi) immunopurification; (vi) aqueous two-phase system; and (vii) aqueous two-phase floatation. A summary of the purification schemes for various bacterial and fungal lipases are also provided.

Received 14 MAY 2014 Revised 05 AUG 2014 Accepted 28 AUG 2014

Keywords: Aqueous two-phase system · Lipase purification · Microbial lipases · Reverse micellar system

1 Introduction The importance of lipases (triacylglycerol acylhydrolases, E.C. 3.1.1.3) in our current world are undeniable and the impact of this group of enzymes is clearly seen in their Correspondence: Dr. Pau Loke Show, Department of Chemical and Environmental Engineering, Faculty of Engineering, University of Nottingham, Jalan Broga, 43500 Semenyih, Selangor Darul Ehsan, Malaysia Email: [email protected] Abbreviations: ATPF, aqueous two-phase floatation; ATPS, aquaeous twophase system; CTAB, cetyltrimethylammonium bromide; DEAE, diethylaminoethyl; EOPO, an unsystematic copolymer of 50% ethylene oxide and 50% propylene oxide; IL, ionic liquid, LCST, lower critical solution temperature; NHS, N-hydroxylsuccin-imide; PANCMA, poly(acrylonitrile-co-maleic acid; PAG, poly(α-allyl glucoside); PEG, polyethylene glycol; PPMM, polypropylene microfiltration membrane; PSLG, poly(γ-stearyl L-glutamate); RMS, reverse micellar system; TX-114,Triton X-114

© 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

widespread application in many industries including organic synthesis, paper manufacturing, oleochemistry, dairy, cosmetics, perfume, biosensors, and detergents [1]. Lipases catalyze the hydrolysis of fats into fatty acids and glycerol at the water-lipid boundary, as well as the synthesis of esters from fatty acids and glycerol at the waterinsoluble substrate boundary. The unique capability to react only at the interface between aqueous and nonaqueous phases distinguishes lipases from esterases [2]. Lipases often have high chemo-, region- and enantioselectivity, enabling lipases to be considered as one of the most important industrially utilized biocatalysts [3]. The current trend in lipase research is to focus on microbial lipases rather than lipases derived from animals and plants, as lipases from micro-organisms have the advantages including the ability to catalyse diverse reactions, produce high yields, and reduced production costs. In addition micro-organisms have the advantage of rela-

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Biotechnol. J. 2015, 10 www.biotecvisions.com


tive ease of genetic manipulation. Other major advantages are that microbial lipases are: (i) structurally stable in organic solvents; (ii) independent of cofactors; (iii) catalyze reactions utilizing a wide range of substrates; and (iv) have high enantio-selectivity [4]. A detailed description of bacterial and fungal lipases, as well as a brief account of lipase purification techniques and screening methods, can be found in a recent review by Nagarajan [5]. According to a study conducted by the Freedonia Group Inc. in 2003 [6], the world enzyme industry is worth $5.8 billion US dollars in 2010, and is forecasted to rise 6.8% annually to $8 billion US dollars in 2015. The study covers the total sales of enzymes (including carbohydrases, proteases, polymerases, nucleases. and lipases) for use in various markets such as biotechnology, food, and pharmaceuticals [6]. Saxena et al. [7] pointed out three important factors that drive the growth of the world enzyme market. Firstly, novel advances made in production biochemistry, fermentation processes, and recovery methods have resulted in affordable large scale production of enzymes. Secondly, the development of various applications employing enzymes have greatly increased market demand. Thirdly, enzymes are capable of catalyzing many different reactions, enhancing their collective commercial usefulness [7]. Despite increased growth in world demand for enzymes, only approximately 200 out of 4000 types of enzymes are commercially available on the global market. Of these 200 enzymes, about 20 types are actively mass produced [8]. The various difficulties impeding the industrialization of enzymes are: (i) it is currently difficult to find enzymes with high yield, high activity, and high stability [9]; (ii) enzymes used for drug and food purposes must go through multiple phases of clinical trials and government approval, e.g. U.S. Food and Drug Administration (FDA) which is both time-consuming and costly [10]; and (iii) enzymes for other purposes such as detergents require strict testing and approval by government sectors dealing with environmental safety, e.g. U.S. Environmental Protection Agency (EPA) [11]. This review serves to highlight the recent (2003-2013) advances in microbial lipase purification, including conventional techniques such as ammonium sulphate fractionation, ion-exchange, gel filtration and affinity chromatography, as well as novel techniques such as reverse micellar system (RMS), membrane processes, immunopurification, aqueous two-phase system (ATPS), and aqueous two-phase floatation (ATPF).

2 Conventional lipase purification methods For a detailed review of both conventional and novel methods of lipase purification established prior to the year 2003, please refer to Saxena et al. [12] and Hasan et al. [2].

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2.1 Lipase prepurification steps Commonly microorganisms secrete lipases into the surrounding liquid matrix. These extracellular lipases are normally produced by fermentation. After the removal of biomass and insoluble particles by filtration or centrifugation, the supernatant is concentrated using ultrafiltration, extraction with organic solvents, or precipitation. Roughly 80% of all the purification methods in use include a precipitation step. 60% of purification procedures used ammonium sulphate while 35% used ethanol, acetone or an acid (usually hydrochloric acid) for precipitation of the enzyme. In simple purification processes the precipitation step is usually done in the early stages of recovery, followed by chromatographic separation. Precipitation techniques normally have high yield (87%) compared to chromatography methods which result in lower yields of around 60–70% [13].

2.2 Chromatographic steps for lipase purification Often the required lipase purity cannot be obtained in a single chromatography step. Therefore a series or combination of chromatography steps is required. There are three types of chromatography methods: (i) ion exchange chromatography is the most frequently used technique, with diethylaminoethyl (DEAE) group and carboxymethyl as the main anion and cation exchangers respectively; (ii) gel filtration chromatography is the second most commonly used method, since gel filtration is limited by its lower capacity in loading proteins, it is normally employed in the final polishing step; and (iii) affinity chromatography, where hydrophobic interaction chromatography is the most popular. The common hydrophobic adsorbents are phenyl and octyl functional groups. Another type of affinity chromatography is adsorption chromatography and the most popular adsorbent is hydroxyapatite. Hydrophobic interaction methods are employed due to the large hydrophobic surface around the active sites of lipases. Although expensive, affinity chromatography techniques can reach a purification factor in the range of 2 to 10 for each step [12]. These chromatography methods are under constant development in the search for breakthroughs. For instance, lipase B from Candida antarctica was purified using biomimetic affinity chromatography which involved attaching synthetic ligands onto Sepharose 4B [14]. In this study, a ligand denoted A9–14 with a cyclohexylamine active group achieved a one-step lipase recovery of 73%. A9–14 ligand had a maximum binding capacity of 0.4 mg/mL and showed no adverse effects on lipase activity. Unlike natural ligands, A9–14 is highly stable in 1M NaOH solution, presenting the potential for large scale CalB production [14]. A purified lipase from Pseudomonas aeruginosa BN-1 was found to contain a chaperonin at its N-terminal. This





Table 1. Summary of conventional lipase purification techniques from 2003 to 2013.

Microbe Strain

PFa)/PRb) (%)

Purification Scheme

Stable Lipase Tc)/pH

Optimum Lipase Tc)/pH

Expression Level

MWd) Reff) e) (kDa)/pI

BACTERIA PPTg),

Acinetobacter sp. EH28

80% Ammonium sulphate ultrafiltration and Phenyl-Sepharose hydrophobic interaction chromatography

24.2/47%

Up to 60°C/8–11

50°C/10

57.1 U/ml

–

[58]

Amycolatopsis mediterranei DSM 43304

40% Ammonium sulphate PPTg), Q Sepharose HP column chromatography and Toyopearl Phenyl-650 column chromatography

398/36%

50–60°C/6–9

60°C 8

781.63 U/mg

33

[59]

Aneurinibacillus thermoaerophilus strain HZ

Q-Sephaorse anion exchange chromatography and Sephadex-G75 gel filtration

15.6/19.7%

60–70°C/6–8

65°C/7

43.5 U/mg

50

[60]

Bacillus megaterium AKG-1

50% Ammonium sulphate PPTg)

5.3/22.5%

Up to 50°C/ 5–10.5

55°C/7

414 U/mg

35

[61]

Geobacillus sp train ARM (expressed in E. coli TOP10)

Immobilized metal affinity chromatography

14.6/63.2%

6–9

65°C/8

7092 U/mg

44

[62]

Pseudomonas aeruginosa BN-1

Ultrafiltration, Sephadex G-100 & DEAE Sephadex A50 column chromatography

42.99

Up to 50°C/ 8–9.5

37°C/8

190.46 U/mg

60

[63]

Ralstonia sp. CS274

Ammonium sulphate PPTg), ultrafiltration and Phenyl Sepharose CL-4B column chromatography

3.9/20.8%

Up to 45°C/ 7–10

50–55°C/ 8–9.5

56250 U/mg

28

[64]

Staphylococcus xylosus

Ammonium sulphate PPTg), 37/11.6% heat treatment, 1st Sephycryl S-200, Mono S-Sepharose and 2nd Sephacryl S-200 chromatography

5–8.5

45°C/8.2

1900 U/mg

43

[65]

Streptomyces fradiae var. k11 (expressed in P. pastoris)

HiTrap Q Sepharose XL anion exchange chromatography and Sephacryl S-200 column chromatography

Up to 40°C/ 4–10

55°C/9.8

569 U/mg

28.5

[66]

17.2/33.7%

25–40°C/7–10

45°C/8

187.5 U/mg

60

[67]

24/38.4%

35–45°C/8–10

37°C/9

502 U/mg

27/4.3

[68]

3.4

Up to 20°C/ 4–8.5

35°C/8.5

88.6 U/mg

63.5

[69]

34/42%

10–50°C/8–11

30°C/9

5878.6 U/mg

31.6

[70]

3.7

FUNGI Antrodia cinnamomea B CRC35396

30–70% Ammonium sulphate PPTg) and DEAE-Sepharose chromatography

Aspergillus carneus Ammonium sulphate PPTg) and Octyl-Sepharose column chromatography Aureobasidium pullulans HN2.3

Ammonium sulphate PPTg), Sephadex G75 gel filtration chromatography and DEAESepharose FFh) anion-exchange chromatography

Fusarium solani N4-2

Acetone fractionation, Q-Sepharose anion exchange chromatography


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Table 1. Summary of conventional lipase purification techniques from 2003 to 2013 (continue).

Microbe Strain

Purification Scheme

PFa)/PRb) (%)



Expression Level


FUNGI PPTg),

PPTg),

Penicillium camembertii Thom PG-3

pH ethanol ammonium sulphate PPT and DEAE-cellulose chromatography

22.1/8.7%

–

48°C/6.4

886 U/mg

28.13

[71]

Talaromyces thermophilus

Ammonium sulphate PPTg), Sephacryl S-200 gel filtration and Mono-Q Sepharose anion exchange chromatography

105/29%

Up to 55°C/9–11

50°C/9.5

9808 U/mg

39

[72]

Yarrowia lipolytica (YILip2)

Q-Sepharose FF ion-exchange chromatography and ButylSepharose FFh) hydrophobic interaction chromatography

26.5/23%

25–35°C/4–10

40°C/8

21300 U/mg

38

[73]

a) b) c) d) e) f) g) h)

PF, Purification Factor PR, Protein Recovery T, Temperature MW, Molecular Weight pI, Isoelectric point Ref, Reference PPT, Precipitation FF, Fast Flow

lipase may have potential utilization in novel lipase coupled biotechnology. Its purification method and a summary of microbial lipases purified by conventional methods can be found in Table 1.

3 Novel lipase purification methods 3.1 Reverse micellar system (RMS) A micelle is an agglomeration of surfactant molecules in which the hydrophilic heads of surfactant molecules are in contact with a polar solvent, sequestering the hydrophobic tails in the micelle core. The micelles are microscopically dispersed throughout the polar solvent forming a liquid colloid [15]. A reverse micelle forms in a non-polar solvent, where the hydrophobic tails of the surfactant molecules are in contact with the solvent, sequestering the hydrophilic heads. This is because the exposure of hydrophobic regions to the surrounding solvent is more thermodynamically favorable. Reverse micelles can hold an inner core of water molecules, also known as water pools, enabling the dissolution of hydrophilic organic compounds such as DNA, proteins, and enzymes (Fig. 1). However, the formation of reverse micelles is proportionally decreased with increasing head group charge, due to the thermodynamically unfavorable electrostatic interactions on hydrophilic sequestration [16]. A surfactant is a compound which reduces the surface tension of a liquid, increasing the solubility of organic compounds. The usage of surfactants in RMS is important

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because they expand the interfacial area of the solvent available for increased micelle formation and recovery of enzymes. This results in the acquisition of higher product yields, a greater degree of enzyme purification, and increased specific activity of the recovered enzyme [15]. The overall recovery process consists of two basic steps, a forward extraction process which involves the uptake of proteins from a bulk aqueous solvent into the water pools of reverse micelles in an inorganic solvent, and a back extraction process where the proteins are transferred from the reverse micelles into another aqueous solvent to be recovered. Entrapment of the proteins in

Figure 1. An illustration of a reverse micelle in equilibrium with its surroundings.





Table 2. Summary of RMS purification scheme (2003–2013)

Microbe Strain

Purification Scheme

PFa)/PRb) (%)

Expression Level

MWc) (kDa)/pId)

Refe)

15/80%

> 23 U/mg

18/8.2

[74]

4.09/82.72%

40.28 U/mg

32

[75]

92%

220 U/mg

–

[76]

BACTERIA Pseudomonas sp. CSD3

RMS (50 mM AOT/Isooctane) at 25°C , pH 6.5, 15 min and back extracted with 0.25M NaCl and 15% isopropanol at 25°C/pH 7 for 30 min FUNGI

Aspergillus niger

RMS (Isooctane/butanol/hexanol at 75/15/10 (v/v/v)) with 0.2M CTABg at pH 9 and back extracted with 0.05M potassium phosphate and 1M KCl at 25°C/pH 7

Thermomyces lanuginosa

RMS (100 mM AOT/Isooctane)

a) b) c) d) e)

PF, Purification Factor PR, Protein Recovery MW, Molecular Weight pI, Isoelectric point Ref, Reference

the water pools of reverse micelles prevents protein denaturation upon contact with organic solvents [17]. It is important that the pH of the aqueous phase allows a target protein to maintain a net surface charge opposite to that of the surfactant headgroups, so that the protein is electrically extracted into the top organic phase. However, in back extraction, the pH of the organic phase must be altered to maintain a net surface charge on the protein equal to the surfactant molecules, so the protein can be pushed out from the reverse micelles [18, 19]. The application of RMS is limited for several reasons. The higher the concentration of surfactant, the harder it is to separate and recover the protein; and the range of organic solvents with which the technique operates efficiently are limited by the constraint to avoid protein denaturation while enabling protein solubilization [20]. A summary of microbial lipases purified by RMS from 2003–2013 can be found in Table 2.

3.2 Membrane processes Abreu et al. has successfully purified and immobilized a lipase from Staphylococcus warneri EX17 (SWL) by onestep adsorption onto hydrophobic supports (i.e. Octylsepharose, Immobead 150 and MCI GEL CHP20P) [21]. For all three supports, 90% SWL immobilization was achieved within 3 hours. The immobilized SWL has a molecular mass of 45 kDa and an optimum temperature of 45°C, higher than the 30°C of free SWL, indicating that suitable membranes can increase the pH and thermal tolerance of lipases [22]. SDS-PAGE analysis revealed that MCI GEL CHP20P adsorbed the most protein, but OctylSWL has the best stability in organic solvents (more than 140% residual activity in methanol, ethanol, isopropanol, and n-hexane). Although Immobead-SWL presented a


low 6% esterification of ethyl butyrate after 24 hours, Octyl-SWL and MCI-SWL achieved 28% and 35.6% conversions respectively [21]. In another study, a lipase from Geobacillus sp. SBS-4S was genetically modified to carry an aldehyde tag (Table 3). The benefit of this aldehyde-tagged lipase was that: (i) it did not require any cross-linking reagents such as 2,4,6-trichloro-1,3,5-triazine or glutaraldehyde-activated supports for its purification process; (ii) the aldehyde tag did not influence the catalytic properties of the lipase ; and (iii) the aldehyde-tagged lipase had a higher resistance to temperature than the native lipase (80% and 65% residual activity respectively at 40°C after 20 h) [23]. A novel dual-layer biomimetic membrane was created from the combination of chitosan-tethered poly(acrylonitrile-co-maleic acid) (PANCMA) ultrafiltration hollow fiber coupled with 1-ethyl-3-(dimethylaminoproply) carbodiimide hydrochloride (EDC)/N-hydroxylsuccin-imide (NHS). This dual-layer membrane was successfully used to purify lipases from Candida rugosa in the presence of glutaraldehyde (GA) bound to the membrane [24]. Analysis of the effect of GA concentration on the specific activity of the bound proteins showed that upon an increase of 2–5% GA concentration, the amount of immobilized protein increased from 48.0 to 66.5 mg/m2, and the specific activity increased from 922 to 1123 U/m2. However, a further increase from 5–8% GA concentration saw an incremental increase in immobilized proteins (from 66.5 to 74.2 mg/m2), but a decline in specific activity (from 1123 to 644 U/m2). This was because over-saturation of GA in the presence of chitosan caused the proteins to be immobilized through multiple chemical bonds with both GA and chitosan, reducing their lipolytic activity. Hence, 5% GA was included in both the purification processes for C. rugosa outlined in Table 3 [24].

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Table 3. Summary of membrane purification scheme (2003–2013)

Microbe Strain

Purification Scheme

PFa)/PRb) (%)



Expression Level


BACTERIA Arthobacter sp. (ABL lipase)

Immobilization of lipase onto diethyl amine (DEA) (72/63)ethanolamine (EA) membrane in phosphate buffer at pH 7 and 40°C for 18 h

–

Up to 50°C/ 6–8

37°C/7

20 U/g

–

[77]

Geobacillus sp. SBS-4S

Immobilization of lipase linked with aldehyde tag onto MCFs-NH2 supports for 3 h

–

Up to 40°C

–

307.9 U/mg

–

[23]

Up to 50°C/ 3–7

45°C/7.5

14.3 U/mg

67

[24]

45°C/7

18.7 U/mg

FUNGI Candida rugosa

a) b) c) d) e) f)

Immobilization of lipase on nascent PANCMA membrane at pH 7.5/45°C

33.9%

Immobilization of lipase on dual-layer biomimetic membrane at pH 7/45°C

44.5%

PF, Purification Factor PR, Protein Recovery T, Temperature MW, Molecular Weight pI, Isoelectric point Ref, Reference

Polypropylene microfiltration membrane (PPMM) is a hydrophobic membrane that has potential industrial value because it has well controlled porosity and is chemically inert. Deng et al. (2004) investigated the extent of lipase immobilization on nascent PPMM, poly(α-allyl glucoside) (PAG)-modified PPMM and poly(γ-stearyl L-glutamate) (PSLG)-modified PPMM [25]. The purpose of grafting PAG and PSLG onto PPMM was to increase the biocompatibility of the membrane. Nascent PPMM was found to have the highest protein adsorption (82.7 mg/m2), moderate specific lipase activity (69.9 U/mg), and moderate activity retention (57.5%). The PSLG-modified PPMM (3.6 wt%) has moderate protein adsorption (74.6 mg/m2) as well as the highest specific lipase activity (88.1 U/mg) and activity retention (72.4%). The PAG-modified PPMM (3.7 wt%) has the lowest recorded results, protein adsorption of 62.5 mg/m2, specific lipase activity of 60.7 U/mg, and activity retention of 49.9%. The low values observed for PAG-modified PPMM was because regions of the membrane with high hydrophilicity prevented lipase adsorption. However, PSLG-modified PPMM has greater hydrophobicity than nascent PPMM due to its long stearyl chains scattered across the membrane, hence higher activity retention [25].

3.3 Immunopurification When affinity chromatography is applied to purify a target protein by using an antibody-antigen system, it is known as immunopurification or immunoaffinity chromatogra-

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phy. Immunopurification is a powerful technique due to its high selectivity for specified proteins, having reported protein purification factors of 1000 to 10000 fold in a single step [26]. This technique can be used to perform difficult separations that other techniques are not able to accomplish. Monoclonal antibodies and affinity-purified polyclonal antibodies are most commonly used in immunopurification. The choice of antibody depends on two important factors: (i) the availability of a suitable monoclonal antibody for the target protein; and (ii) the composition and concentration of the contaminants present [27]. Previously immunopurification was regarded as one of the most costly affinity techniques, especially when utilizing monoclonal antibodies. However, due to the development of monoclonal antibody technology and the discovery of cost-effective methods for antibody mass production, the costs of immunopurification has been reduced, resulting in the growth of industrial applications involving monoclonal antibodies [28]. Rahimi and co-workers synthesized and compared the activity of two novel monoclonal antibodies denoted BF11A and VNH9 on the immobilization of Candida rugosa lipase [29]. After formation of an immunocomplex with BF11A and VNH9, the residual activity of the lipase was 99% and 92% respectively, which shows good potential for utilization in purification and immobilization systems [29]. In a separate study, Candida antarctica lipase B (CalB) was genetically engineered to bind with a free cysteine via a histone (His) tag to produce CalB-HisGGC and CalB-





C311A-His. A heterobifuntional NHS-PEG-maleimide was used as a spacer, where PEG is poly(ethylene glycol). A maleimide-activated glass support was used and a lipase purification of 10,000 fold was achieved in a single step [30].

3.4 ATPS The use of conventional organic-aqueous solvent systems in protein purification is not practical owing to the fact that labile proteins would be denatured by the organic solvent. On the other hand, ATPS is well suited for the purifi-

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cation of biomolecules (such as proteins, DNA, RNA, bionanoparticles, organelles, and cells) as it contains a high water content. ATPS shows great promise for the downstream processing of proteins due to advantages such as: potential for scaling up to industrial scale, continuous mode of operations, relative ease to integrate with other purification processes, low toxicity of phase-forming chemicals, and high biocompatibility [31]. Upon mixing of aqueous solutions containing two different incompatible polymers, they will separate into two distinct phases because of steric exclusion. ATPS makes use of this incompatibility for extraction and purification

Figure 2. An illustration of an extractive fermentation process using the recycling ATPS developed by Show et al. [40] (*Reproduced with permission from Elsevier).


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Table 4. Summary of ATPS purification scheme (2003–2013)

Microbe Strain

Purification Scheme

PFa)/PRb) (%)

Expression Level

MWc) (kDa)/pId)

Refe)

BACTERIA Acinetobacter sp. MTCC 6816

ATPS (24% w/v PEG 8000/15.5% w/v potassium phosphate) optimized by RSM at pH 7 and Octyl Sepharose CL-4B

14.92/74.6%

88.8 U/mg

32

[78]

Bacillus sp. ITP-001

80% ammonium sulphate PPT, dialysis and ATPS (25% w/w [C8mim]Cl/30% w/w potassium phosphate) at pH 7/25°C

51

–

54/3

[48]

Burkholderia cenocepacia ST8

4 successive recycling ATPS (EOPO/potassium phosphate)

14/99%

17 U/ml

34.5/6.3

[40]

Burkholderia cepacia

Recycling ATPS extractive fermentation (10% w/w EOPO 3900)

10.4/99.3%

45.4 U/ml

34/6.3

[79]

Burkholderia pseudomallei

ATPS (16% w/w 2-propanol/16% w/w potassium phosphate) and 4.5% w/v NaCl

13.5/99%

–

40.3

[80]

ATPS (PEG 6000/potassium phosphate) and 1% w/w NaCl at pH 7

12.42/93%

–

40.3

[81]

92.1%

15–18.5 U/ml

36.2

[82]

80.4%/5.84

1114.6 U/g

–

[42]

ATPS (9.6% w/w PEG 8000/1% w/w Dextran T500) at pH 9 Trichosporon laibacchii

ATPS (12% w/w PEG 4000/13% w/w potassium phosphate) and 2% NaCl at pH 7 FUNGI

Aspergillus niger NRRL3

ATPS (PEG 4000/Ci) at pH 5.2

Candida antarctica (lipase B)

ATPS (25% w/w [C8mim]Cll/30% w/w potassium phosphate buffer solution) at 25°C and pH 7

a) b) c) d) e)

–

35/4.4

[41]

–

35.3/6

[83]


purposes. Phase separation also occurs between a salt with high ionic strength and a polymer. Since the salt is capable of retaining a huge amount of water, it also displays a phase separation phenomenon. The most popular pair of polymers used in polymer-based ATPS are PEG and dextran, while the commonly used polymer-salt ATPS includes PEG-potassium phosphate ATPS and PEG-magnesium sulphate ATPS [32]. Target proteins can be isolated from other proteins and cell debris using ATPS. Normally the upper phase is more hydrophobic in nature, for example an aqueous phase containing PEG [33]. Successful manipulation of the partition coefficient of the two-phase partitioning system determines the separation of the target proteins from other proteins. The partition coefficient can be changed by: (i) adjusting the average molecular weight of the polymers used; (ii) the type of ions in the aqueous phase; (iii) the ionic strength of the salt; and (iv) by including an extra salt such as NaCl. Alternatively, the polymer can also be modified by attaching hydrophobic groups to change the partitioning of the pro-

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30.5 2.6/95.9%

tein. Since ATPS is a very soft and non-invasive method of protein purification, protein denaturation or loss of biocatalyst activity is rare. The reason for this is due to the protection to the proteins offered by the large water pool and the low interfacial tension. The polymers used also play an important role by acting as stabilizers for the proteins [34]. Conventional ATPS uses phase-partitioning chemicals that are cost ineffective and difficult to recycle, such as PEG, dextran, polyvinyl alcohol (PVA), and ammonium sulphate [35]. For instance, it is theoretically possible to separate PEG from a PEG-(NH4)2SO4 solution, whereby extra ammonium sulphate has to be added to the PEGrich phase until the salt concentration reaches its solubility limit [36]. Alternatively, PEG can also be recovered by thermoseparation, but its cloud point temperature lies about 180°C for low molecular weights and 95°C for those with 200 kDa and above [37]. Hence, research into novel thermoseparating polymers resulted in the development of an unsystematic copolymer of 50% ethylene oxide (EO)





and 50% propylene oxide (PO), named EOPO [38]. Possessing a cloud point temperature as low as 47°C, EOPO can be recycled while showing high protein yields [39]. In a recycling ATPS, there is a primary ATPS where the target protein is extracted into an EOPO-rich top phase; and a secondary ATPS where the EOPO-rich top phase from the primary ATPS is transferred and heated to its lower critical solution temperature (LCST), separating into two phases: a top water phase entrapping the target protein, and a bottom phase of concentrated EOPO solution [39]. The concentrated EOPO solution from the secondary ATPS can be reused to further extract proteins from the salt solution in the primary ATPS (Fig. 2) [40]. ATPS (PEG 4000/Ci) has been used to extract lipases from Aspergillus niger NRRL3 from solid culture media with a high purification factor. This methodology offers several advantages: (i) recovery of enzyme in a polymerpoor medium; (ii) the citrate buffer generates less pollution than phosphate; and (iii) the solid culture medium is easy to reproduce [41]. The details of the purification procedure can be found in Table 4. A recent study successfully combined ATPS with enzyme immobilization to enhance lipase recovery. Diatomites were utilized as the lipase immobilization support in the top phase of ATPS (12% w/w PEG 4000/13% w/w potassium phosphate). This led to a high yield and direct single step purification of the lipase, discarding the need for an extractive bottom phase. The details of the purification scheme is shown in Table 4 [42]. Apart from the polymer-salt ATPSs, ionic liquid (IL)-based ATPSs has received much attention in the past decade owing to the unique features of ILs such as nonflammability and high chemical stability [43, 44]. In addition, the properties of ILs can often be rationally tuned by selecting different combination of ions, earning ILs their “designer solvents” designation [45]. ILs and inorganic salts-based ATPS have been reported to exhibit higher protein recovery than conventional polymer-salt systems [46–48]. In one study an ATPS was formed from pairing an imidazolium based IL with an alkylsulfate anion to extract lipase A from Candida antarctica. It was found that 1-ethyl-3-methylimidazolium butyl sulphate [C2MIM][C4SO4] coupled with ammonium sulphate achieved an enzyme recovery of 99%. Lipase A has an optimum temperature above 90°C, higher than that of lipase B from the same fungus species [49]. The advantages of this method are: (i) short processing time (~ one hour); (ii) high enzyme recovery; (iii) low heat utility requirement; (iv) mild operating conditions; and (v) ILs as the medium for biocatalytic transformations [49]. Extractive fermentation using ATPS is another novel technique that allows continuous separation of products during the fermentation stage. Advantages of this system include: (i) high mass transfer due to low interfacial tension; (ii) highly selective product separation; (iii) high



yield of biomolecules (iv) ease of continuous operation; (v) biocompatibility; (vi) suitability for systems with product inhibition; and (vii) ecofriendlyness. This system, also known as ATPS-based in situ product recovery, allows simultaneous product synthesis and removal [50]. In extractive fermentation using ATPS the system is manipulated in such a way that the microbial cells are immobilized in one of the ATPS phases, while the required biomolecules partitioned to the other phase [50]. Mostly this system has been used to purify bacteria lipases due to: (i) the high yield of products from bacterial strains with short doubling times; (ii) the inability of bacterial strains to metabolize polymers such as PEG or EOPO under normal culture conditions (temperature below 47°C); (iii) the ease of genetic manipulation of bacterial genomes; (iv) negligible alterations to the physical properties, such as viscosity and rheology, of the system after addition of bacteria, (unlike fungal fermentation); and (v) no mycelia formation which disturbs phase separation and product recovery. A list of extractive fermentations using ATPS for various bacteria and fungi has been outlined by Banik et al. (2003) [50]. A summary of microbial lipases purified by ATPS from 2003–2013 can be found in Table 4.

3.5 Aqueous two-phase flotation (ATPF) Aqueous two-phase flotation (ATPF) is a novel recovery method which integrates ATPS with solvent sublation (SS) [51]. SS is an adsorptive bubble separation technique which works by bubbling nitrogen gas through the bottom enzyme-rich solution, capturing surface-active (or hydrophobic) compounds by adsorption onto the nitrogen gas, and the ascending bubbles then dissolve in the top polymer phase. The rest of the system is identical to ATPS and hence this purification method inherits the advantages of ATPS [52, 53]. Similar to recycling ATPS, recycling ATPF also has two steps, a primary ATPF where the target protein is extracted by a top EOPO phase; and a secondary ATPF formed from the top phase of the primary ATPF, where it thermoseparates at its LCST into a top aqueous phase containing the target protein and a bottom phase comprising of concentrated EOPO solution. The concentrated EOPO solution is reused to further purify the desired protein in the crude feedstock via the primary ATPF (Fig. 3) [53]. A recycling ATPF (50% (w/w) EOPO 3900/250 g/L ammonium sulphate) has been utilized to extract Burkholderia cepacia ST8 lipase directly from the culture broth in a single recycling step. The optimized conditions of this system were 40% (w/w) crude feedstock, 30 ml/min nitrogen flow rate at 60 min flotation time, and pH 6. To induce thermoseparation, the EOPO phase was removed from the primary extraction and submerged in a 65°C water bath for 15 minutes. A purification of 17.75 fold and protein recovery of 98.22% were achieved [53].

9




Figure 3. An illustration of the recycling ATPF process developed by Show et al. [53] (*Reproduced with permission from Elsevier).

B. cepacia ST8 lipase was also successfully extracted using an alcohol-salt ATPF, in this case 50% (w/w) 2-propanol/250g/L potassium phosphate. The best experiment was conducted at pH 8.5 with a nitrogen flow rate of 30 ml/min at a flotation time of 30 minutes. The lipase was purified 14.4 fold and the protein recovery was 99.2%. The average sustainable recovery of 2-propanol and potassium phosphate were shown to be up to 70% and 61% respectively [53]. A summary of microbial lipases

purified by ATPF between 2003–2013 can be found in Table 5.

3.6 Aqueous micellar two-phase system (AMTPS) Aqueous micellar two-phase system (AMTPS) is another variation on ATPS which utilizes a binary phase micellar system with the help of surfactants at low concentrations [54]. Micelles are induced in aqueous solutions when the

Table 5. Summary of ATPF purification scheme for Burkholderia cenocepacia ST8 (2003–2013)

Microbe Strain

Purification Scheme

PFa)/PRb) (%)

Expression Level

MWc) (kDa)/pId)

Refe)

BACTERIA Burkholderia cenocepacia ST8

Recycling ATPF (50% w/w EOPO 3900/250 g/L ammonium sulphate) at 15 min heating at 65°C/pH 6

12.26/98.81%

111.15 U/ml

33

[53]

Burkholderia cenocepacia ST8

Recycling ATPF (50% w/w 2-propanol/250g/L potassium phosphate)/pH 8.5

14.4/99.2%

–

34

[79, 84]

a) b) c) d) e)

10







concentration of surfactants reaches a minimum limit known as critical micelle concentration (CMC) [55]. Once the temperature of the surfactant solution reaches a minimum limit, called cloud-point temperature, two coexisting phases are formed in AMTPS [56]. When the temperature is above the cloud-point, the surfactant molecules self-aggregate to maintain an aqueous (surfactant-depleted) phase and a coacervate (micelle-enriched) phase [55]. The most common phase-forming surfactants used are Triton X-114 (polyoxyethylene detergent, C14EO6) and other alkyl polyoxyethylenes (CmEOn) such as n-decyltetraethylene oxide (C10EO4). A study of the effects of the surfactants Triton X-114 (TX-114) and Pluronic (L31, L61, L81, and L121) on the extraction of Burkholderia sp. ST8 was conducted (Fig. 4). It was found that across a concentration range of 5–20% (w/w), Pluronic L121 has the lowest average cloud point of 10.3°C, whereas TX-114 at 20% (w/w) has the highest cloud point of 37°C. Lipase recovery was obtained only with Pluronic L81 and TX-114, where the better surfactant is 20% (w/w) Pluronic L81, achieving a partition coefficient (KL) of 0.032. From further testing it was concluded that 24% (w/w) Pluronic L81 with 0.5% (w/w) KCl achieved the highest lipase recovery with a selectivity of 0.035 and purification factor of 7.2. The lipase has a molecular weight of 36.7 kDa [57].

4 Concluding remarks The numerous studies regarding the identification and purification of lipases from micro-organisms have highlighted the potential of microbial lipases for industrial applications. Most of the reported works purify lipases in a multistep manner. However, the multistep purification scheme comes at a cost when applied at an industrial scale, as several types of specific equipment are required. The size of the equipment depends on the quantity and purity of the target – the higher the quantity and purity, higher the cost of equipment. Another factor to be taken into account is the raw materials needed, for instance the source of micro-organisms, the biomass used, the nutrients supplied, and other chemicals. On the other hand, there are recent developments where both conventional and novel techniques have been shown to achieve a high yield and purification factor in a single step, such as creating synthetic ligands to increase the yield of one-step biomimetic affinity chromatography [14], and the use of a heterobifuntional NHS-PEGmaleimide spacer [30]. Methods that recycle chemicals for purification purposes will in no doubt become trendsetters, with an example of such a setup being the inclusion of the thermoseparating polymer EOPO in ATPF [53]. These techniques hold, as yet, unrealized potential, and once matured will pave the way for technologies that will be employed in future industries.


Figure 4. An illustration of the AMTPS process developed by Ooi et al. [57] (*Reproduced with permission from Elsevier).

An overview of the works cited revealed that there is no one optimum condition to cultivate micro-organisms and extract the corresponding lipases. Culture conditions vary from one microbe strain to another, leading to the combination of different techniques to achieve the best protein yield and lipase homogeneity. Regardless, methods such as membrane technology and immunoaffinity chromatography are a promising field of research, which should be exploited for potential commercial applications.

This work is supported financially by SATU Joint Research Scheme (RU022E-2014) from University of Malaya, Fundamental Research Grant Scheme (FRGS/1/ 2013/SG05/UNIM/02/1), (FP005-2013B), and Ministry of Science, Technology and Innovation Grant (MOSTI-0202-12-SF0256) from Malaysia. The authors declare no financial or commercial conflict of interest.

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5 References Tau Chuan Ling is a Professor at the [1] Aravindan, R., Anbumathi, P., Viruthagiri, T., Lipase applications in food industry. Indian J. Biotechnol. 2007, 6, 141–158. [2] Hasan, F., Shah, A. A., Hameed, A., Industrial applications of microbial lipases. Enzyme Microb. Tech. 2006, 39, 235–251. [3] Gotor-Fernández, V., Brieva, R., Gotor, V., Lipases: Useful biocatalysts for the preparation of pharmaceuticals. J. Mol. Catal. B-Enzym. 2006, 40, 111–120. [4] Jaeger, K.-E., Reetz, M. T., Microbial lipases form versatile tools for biotechnology. Trends biotechnol. 1998, 16, 396–403. [5] Nagarajan, S., New Tools for Exploring “Old Friends – Microbial Lipases”. Appl. Biochem. Biotech. 2012, 168, 1163–1196. [6] The Freedonia Group Inc., World Enzymes to 2015 – Demand and Sales Forecasts, Market Share, Market Size, Market Leaders, Ohio, USA 2014, pp. 1–338. [7] Sharma, R., Chisti, Y., Banerjee, U. C., Production, purification, characterization, and applications of lipases. Biotechnol. Adv. 2001, 19, 627–662. [8] Li, S., Yang, X., Yang, S., Zhu, M., Wang, X., Technology Prospecting on Enzymes: Application, Marketing and Engineering. Comput. Struct. Biotechnol. J. 2012, 2, 1–11. [9] Shuler, M. L., Kargi, F., Bioprocess Engineering, Prentice Hall New York 2002. [10] How Drugs are Developed and Approved. U.S. Food and Drug Administration, 2013. [11] U.S. Environmental Protection Agency, in: Federal Register, 2013, 76, pp. 55264–55268. [12] Saxena, R. K., Sheoran, A., Giri, B., Davidson, W., Purification strategies for microbial lipases. J. Microbiol. Methods 2003a, 52, 1–18. [13] Aires-Barros, M. R., Taipa, M. A., Cabral, J. M. S., Isolation and purification of lipases.in Wolley P., Petersen S.B. (Eds), Lipases: Their Structure, Biochemistry and Application, Cambridge University Press (UK) 1994, pp 234–270. [14] Yao, H., Zhang, T., Xue, H., Tang, K., Li, R., Biomimetic affinity purification of Candida antarctica lipase B. J. Chromatogr. B 2011, 879, 3896–3900. [15] Levashov, A. V., Klyachko, N. L., Reverse micellar systems, Enzymes in Nonaqueous Solvents, Springer 2001, pp. 575–586. [16] Shin, Y. O., Vera, J. H., Solubilization limit of lysozyme into DODMAC reverse micelles. Biotechnol. Bioeng. 2002, 80, 537–543. [17] Yu, Y.-c., Chu, Y., Ji, J.-Y., Study of the factors affecting the forward and back extraction of yeast-lipase and its activity by reverse micelles. J. Colloid Interface Sci. 2003, 267, 60–64. [18] Dekker, M., Hilhorst, R., Laane, C., Isolating enzymes by reversed micelles. Anal. Biochem. 1989, 178, 217–226. [19] Pessoa Jr, A., Vitolo, M., Recovery of inulinase using BDBAC reversed micelles. Process Biochem. 1998, 33, 291–297. [20] Basheer, S. A., Thenmozhi, M., Reverse Micellar Separation of Lipases: A Critical Review. Int. J. Chem. Sci. 2010, 8, 57–67. [21] de Abreu, L., Fernandez-Lafuente, R., Rodrigues, R. C., Volpato, G., Ayub, M. A. Z., Efficient purification-immobilization of an organic solvent-tolerant lipase from Staphylococcus warneri EX17 on porous styrene-divinylbenzene beads. J. Mol. Catal. B: Enzym. 2014, 99, 51–55. [22] Volpato, G., Filice, M., De Las Rivas, B., Rodrigues, R. C., et al. Purification, immobilization, and characterization of a specific lipase from Staphylococcus warneri EX17 by enzyme fractionating via adsorption on different hydrophobic supports. Biotechnol. Progr. 2011, 27, 717–723. [23] Wang, A., Du, F., Wang, F., Shen, Y., et al., Convenient one-step purification and immobilization of lipase using a genetically encoded aldehyde tag. Biochem. Eng. J. 2013, 73, 86–92.

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Institute of Biological Sciences, University of Malaya. He is a Chartered Engineer of the Engineering Council, United Kingdom (CEng) and has been a member of the Institution of Chemical Engineers (IChemE) since 2011. He received his BSc in Biotechnology and MSc in Environmental Engineering from the Universiti Putra Malaysia, and his PhD in Chemical Engineering from Birmingham University, United Kingdom. He has more than 10 years of research experience in downstream processing and bioprocess engineering. To date he has published more than 100 ISI cited articles in international journals and presented more than 50 conference papers.

Pau Loke Show is Assistant Professor at the Department of Chemical and Environmental Engineering at The University of Nottingham, Malaysia. He received his PhD in Bio-Process Engineering from the Universiti Putra Malaysia. He is an associate member for the Institution of Chemical Engineers (IChemE). His research focuses on bioprocess engineering from upstream to downstream processing in biotechnology and industrial microbiology. He is also interested in fermentation technology using green methods to produce sustainable chemistry. In the last 3 years, he has published 15 SCI journal articles and presented more than 10 conference papers. In addition, he is currently serving on the international scientific committees of several international conferences and also serves as reviewer for more than 20 peer-reviewed journals. He has also established collaborations with well-known international researchers from countries including the United Kingdom, Taiwan, New Zealand, throughout his career.

Chung-Hong Tan is currently a PhD student, studying in the University of Nottingham in Malaysia supported by a university scholarship. In the same institution, he pursued an honours degree in Chemical Engineering. During his undergraduate years, he had experience in treating palm oil mill effluent (POME) with flocculation. He would like to thank the following supervisors for their continued support: Prof. Chang Jo-Shu, Dr. Chen Chun-Yen, Prof. Ling Tau Chuan, Dr. Lam Hon Loong and Dr. Show Pau Loke.





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