Microbial Cell Factories
`
`BioMed Central
`
`Open Access
`Review
`Strategies for the recovery of active proteins through refolding of
`bacterial inclusion body proteins
`Luis Felipe Vallejo and Ursula Rinas*
`
`Address: Biochemical Engineering Division, GBF German Research Center for Biotechnology, Mascheroder Weg 1, 38124 Braunschweig, Germany
`
`Email: Luis Felipe Vallejo - lfv@gbf.de; Ursula Rinas* - uri@gbf.de
`* Corresponding author
`
`Published: 02 September 2004
`
`Microbial Cell Factories 2004, 3:11
`
`doi:10.1186/1475-2859-3-11
`
`Received: 29 June 2004
`Accepted: 02 September 2004
`
`This article is available from: http://www.microbialcellfactories.com/content/3/1/11
`
`© 2004 Vallejo and Rinas; licensee BioMed Central Ltd.
`This is an open-access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0),
`which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
`
`Abstract
`Recent advances in generating active proteins through refolding of bacterial inclusion body proteins
`are summarized in conjunction with a short overview on inclusion body isolation and solubilization
`procedures. In particular, the pros and cons of well-established robust refolding techniques such as
`direct dilution as well as less common ones such as diafiltration or chromatographic processes
`including size exclusion chromatography, matrix- or affinity-based techniques and hydrophobic
`interaction chromatography are discussed. Moreover, the effect of physical variables (temperature
`and pressure) as well as the presence of buffer additives on the refolding process is elucidated. In
`particular, the impact of protein stabilizing or destabilizing low- and high-molecular weight additives
`as well as micellar and liposomal systems on protein refolding is illustrated. Also, techniques
`mimicking the principles encountered during in vivo folding such as processes based on natural and
`artificial chaperones and propeptide-assisted protein refolding are presented. Moreover, the
`special requirements for the generation of disulfide bonded proteins and the specific problems and
`solutions, which arise during process integration are discussed. Finally, the different strategies are
`examined regarding their applicability for large-scale production processes or high-throughput
`screening procedures.
`
`Background
`Recombinant DNA technology made available several
`simple techniques for transferring and efficiently express-
`ing desired genes in a foreign cell. Thus, it was thought
`that unlimited and inexpensive sources of otherwise rare
`proteins would become accessible. It soon was observed
`that the host cell had a great influence on the quality and
`quantity of the produced recombinant protein. For exam-
`ple, recombinant protein production in mammalian cells
`yields a biologically active protein with all the required
`posttranslational modifications. However, mammalian
`cell cultivation is characterized by low volumetric yields
`of the recombinant protein, long cultivation times and
`
`requirements for expensive bioreactors and medium com-
`ponents. All these points have a great impact on the pro-
`duction costs. On the other hand, bacterial cultivation
`processes are based on inexpensive media in which fast
`growth and high cell concentrations can be obtained.
`These high cell concentrations combined with higher pro-
`duction rates of the bacterial expression system result in
`higher volumetric productivities. However, the produc-
`tion of recombinant proteins in bacteria such as
`Escherichia coli frequently yields an inactive protein, aggre-
`gated in the form of so-called inclusion bodies.
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`Though, producing an inactive target protein in the form
`of inclusion bodies is an important drawback, it also has
`several advantages such as the high degree of purity of the
`target protein in the aggregate fraction and the increased
`protection from proteolytic degradation compared to the
`soluble counterpart. Inclusion bodies have long been con-
`sidered completely inert towards in vivo dissolution; only
`recently it was shown that proteins can be resolubilized in
`vivo from inclusion body deposits [1]. Although inclusion
`bodies in general consist of inactive proteins, E. coli can be
`the superior expression system compared to eukaryotic
`expression systems when the activity of the recombinant
`protein can be regained through refolding from the pro-
`duced inclusion bodies. However, one needs to consider
`that the decision to select a specific expression system fre-
`quently is based on more trivial reasons such as staff
`knowledge and available equipment and facilities of the
`producing company/institute.
`
`A good example to demonstrate the diverse routes that
`can be used for recombinant protein production is the
`manufacturing of tissue-type plasminogen activator (tPA).
`This protein enables the dissolution of blood clots and is
`used therapeutically for the treatment of myocardial inf-
`arction, thrombosis, pulmonary embolism, and strokes.
`To assure sufficient tPA for such a widespread application,
`an economic production process is a necessity. From the
`beginning, both the mammalian as well as the microbial
`route were explored for the production of tPA [2]. tPA is a
`fairly large (527 amino acids) monomeric protein con-
`taining 17 disulfide bridges. Because of this complexity,
`tPA was first produced in E. coli in the form of inclusion
`bodies while the mammalian expression system yielded
`an active protein that was secreted into the culture
`medium. More recently, obtaining active tPA through
`secretion into the periplasm of E. coli was attempted [3-5].
`The early unsatisfactory yields have been improved [6,7]
`rendering the E. coli secretion system as a future potential
`alternative route to generate functional tPA. Other recom-
`binant organisms such as yeast [8], fungi [9] or insect cells
`[10] have not yet been considered as industrial producers
`for this protein.
`
`Initially, the recombinant tPA introduced into the market
`was obtained from genetically engineered mammalian
`cells [2]. At that time, generating biologically active tPA
`from E. coli produced material was a process with a poor
`overall yield [2]. Today, the majority of commercial tPA
`(alteplase, Activase®) is still produced using a mammalian
`expression system (Genentech: http://www.gene.com/
`gene/products/information/cardiovascular/activase/). In
`addition, an amino substituted tPA produced by the
`mammalian expression system with increased half-life
`(tenecteplase) was developed. Alternatively, a non-glyco-
`sylated, truncated tPA (reteplase, Retavase®) produced in
`
`E. coli in form of inclusion bodies and afterwards refolded
`to its biologically active form is now on the market (Cen-
`tocor: http://www.retavase.com/) and apparently gains
`market share at the cost of the mammalian-derived prod-
`uct(s) (see Genentech 2004 First Quarter Report).
`
`Thus, continuous research effort focused on developing
`new refolding techniques or improving existing ones by
`including novel refolding aiding agents can make the bac-
`terial inclusion body system an excellent alternative to the
`mammalian expression system or other expression sys-
`tems that can directly generate active proteins with a com-
`plex disulfide bond structure. The foremost aim in
`improving protein refolding from E. coli produced inclu-
`sion bodies is to increase both the allowed protein con-
`centrations during the refolding process and the final
`refolding yield. Recent advances in this area are summa-
`rized in conjunction with a short overview on inclusion
`body isolation and solubilization procedures. Moreover,
`the different techniques are discussed regarding their
`applicability for large-scale production processes or high-
`throughput screening procedures.
`
`Isolation and solubilization of inclusion bodies
`A high degree of purification of the recombinant protein
`can be achieved by inclusion body isolation [for recent
`reviews on various aspects of inclusion body formation
`and renaturation of inclusion body proteins please refer
`also to [11-18]]. Inclusion bodies are in general recovered
`by low speed centrifugation of bacterial cells mechanically
`disrupted either by using ultrasonication for small, French
`press for medium, or high pressure homogenization for
`large scale. Main protein contaminants in the crude inclu-
`sion body fraction are proteins from the cell envelope, the
`outer membrane proteins [19]. These proteins are not
`integral inclusion body contaminants but coprecipitate
`together with other insoluble cell material during inclu-
`sion body recovery. Lysozyme-EDTA treatment before cell
`homogenization facilitates cell disruption. Addition of
`detergents such as Triton X-100 and/or low concentra-
`tions of chaotropic compounds either prior to mechanical
`cell breakage or for washing crude inclusion body prepa-
`rations allow the removal of membrane proteins or other
`nonspecifically adsorbed cell material [11-14].
`
`After their isolation, inclusion bodies are commonly sol-
`ubilized by high concentrations of chaotropic agents such
`as guanidinium hydrochloride or urea. Although expen-
`sive, guanidinium hydrochloride is in general preferred
`due to its superior chaotropic properties. Moreover, urea
`solutions may contain and spontaneously produce
`cyanate [20], which can carbamylate the amino groups of
`the protein [21]. In addition, inclusion body solubiliza-
`tion by urea is pH dependent and optimum pH condi-
`tions must be determined for each protein [22]. There are
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`also reports that inclusion bodies can be solubilized at
`extreme pH in the presence or absence of low concentra-
`tions of denaturants [23-25]. However, extreme pH treat-
`ments can result in irreversible protein modifications such
`as deamidation and alkaline desulfuration of cysteine res-
`idues [26]. Finally, inclusion bodies can be solubilized
`with different types of detergents [27,28], low concentra-
`tions of denaturants [29,30], or even by utilization of the
`aggregation suppressor arginine [29]. Inclusion body pro-
`teins solubilized under these mild conditions can possess
`a native-like secondary structure [28-30], and may even
`reveal some biological activity [29,31]. It has also been
`demonstrated that the utilization of milder solubilization
`conditions can lead to higher final refolding yields com-
`pared to solubilization by high concentrations of gua-
`nidinium hydrochloride or urea [27].
`
`In addition to the solubilizing agent, the presence of low
`molecular weight thiol reagents such as dithiothreitol
`(DTT) or 2-mercaptoethanol is generally required. These
`substances will reduce nonnative inter- and intramolecu-
`lar disulfide bonds possibly formed by air oxidation dur-
`ing cell disruption and will also keep the cysteines in their
`reduced state [14,15]. Optimum conditions for disrup-
`tion of existing disulfide bonds are found at mild alkaline
`pH since the nucleophilic attack on the disulfide bond is
`carried out by the thiolate anion. Residual concentrations
`of reducing substances can negatively affect the refolding
`process, thus, they are frequently removed (e.g. by dialy-
`sis) before starting the refolding procedure. As an alterna-
`tive,
`immobilized
`reducing
`agents
`(e.g. DTT;
`VectraPrime™, Biovectra) could simplify reducing agent
`removal by centrifugation after the solubilization process.
`Finally, the pH must be reduced before the removal of the
`reducing agent from the solution containing the solubi-
`lized protein to prevent the formation of undesired
`disulfide bonds.
`
`Principles of refolding solubilized and unfolded proteins
`Correct refolding versus aggregation
`In general, the methods used for inclusion body solubili-
`zation result in a soluble protein that is devoid of its
`native conformation. This protein must then be trans-
`ferred into conditions that allow the formation of the
`native structure (e.g. low denaturant concentration).
`Moreover, appropriate redox conditions have to be estab-
`lished when the protein contains disulfide bonds in the
`native state. When proper conditions for refolding are
`identified, the refolding process can require a few seconds
`or several days. During this period, the correct refolding
`pathway competes, often in disadvantage, with misfold-
`ing and aggregation of the target protein (Figure 1). Pro-
`tein refolding involves intramolecular interactions and
`follows first order kinetics [32-35]. Protein aggregation,
`however, involves intermolecular interactions and, thus,
`
`Simplified model of correct folding versus misfolding and aggregationFigure 1
`
`
`Simplified model of correct folding versus misfolding and
`aggregation. The correct protein folding pathway (1) often
`competes with misfolding (2) and aggregation (3). Aggrega-
`tion occurs among intermediates with exposed hydrophobic
`patches, which are buried in the correctly folded protein
`(blue lines, hydrophilic solvent-exposed parts of the protein;
`red lines: hydrophobic patches).
`
`is a kinetic process of second or higher order, which is
`favored at high protein concentrations [32-35]. In fact,
`refolding yields commonly decrease with increasing ini-
`tial concentrations of the unfolded protein independent
`of the refolding method applied [35-40].
`
`Aggregates are formed by nonnative intermolecular
`hydrophobic interactions between protein folding inter-
`mediates, which have not yet buried their hydrophobic
`amino acid stretches (Figure 1). When the refolding proc-
`ess is beyond these aggregation-prone intermediates, the
`productive folding pathway is favored and aggregation
`does not occur. Therefore, prevention of hydrophobic
`intermolecular interaction during the first steps of refold-
`ing is crucial to allow successful renaturation at high pro-
`tein concentrations. Only recently a non-empirical
`method for predicting the fate of proteins during the
`refolding process was proposed [41]. It is based on the
`second viral coefficient, which indicates the magnitude of
`protein interaction under certain refolding conditions,
`and thus its tendency to aggregate. However, though
`being soluble in the refolding buffer is essential for a pro-
`tein molecule to refold, it does not ensure that it will fold
`into the native form.
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`Are further purification steps required after solubilization of inclusion
`bodies?
`The recombinant target protein represents in general the
`major fraction of the inclusion body proteins. Therefore,
`refolding attempts can be undertaken directly after solubi-
`lization of the inclusion bodies. Some reports, however,
`claim higher refolding yields when the solubilized inclu-
`sion body proteins are purified prior to the refolding
`attempt [36,39,42,43]. Additional purification has been
`recommended when the protein of interest represents less
`than 2–5% of the total cell protein [26] or less than 2/3 of
`the total inclusion body protein [42]. The type of contam-
`inants can also be crucial for the success of the refolding
`process. For example, typical non-proteinaceous contam-
`inants of inclusion body preparations did not affect
`refolding yields of lysozyme, while proteinaceous con-
`taminants, which have a high tendency towards aggrega-
`tion significantly reduced refolding yields [42]. Further
`purification prior to the refolding attempt does not seem
`to be required, even at low target protein concentrations,
`when the solubilized inclusion body proteins are sub-
`jected to refolding conditions during size exclusion chro-
`matography where refolding and purification can occur
`simultaneously [44]. All pros and cons of any further puri-
`fication step have to be carefully evaluated as they cause
`potential protein loss and additional production costs.
`
`Techniques for protein refolding
`Direct dilution
`The simplest refolding procedure is to dilute the concen-
`trated protein-denaturant solution into a refolding buffer
`that allows the formation of the native structure of the
`protein. Most frequently, the final protein concentration
`after dilution is in the 1–10 µg/ml range in order to favor
`the productive refolding instead of the unproductive
`aggregation pathway. Though ideal at laboratory scale,
`this technique has serious drawbacks during scale-up as
`huge refolding vessels and additional cost-intensive con-
`centration steps are required after renaturation.
`
`A major improvement of this technique was the develop-
`ment of a method where the solubilized, denatured pro-
`tein is added in pulses or continuously into the refolding
`buffer [37,40,45-47]. This technique still keeps the sim-
`plicity of the direct dilution method while considerably
`increasing the final concentration of the refolded protein.
`Prerequisite is an appropriate knowledge of the folding
`kinetics of the target protein. The addition of the concen-
`trated protein-denaturant solution should occur at rates
`slower than the rate-determining folding step of the target
`protein, thereby avoiding the accumulation of aggrega-
`tion-prone folding intermediates [37,46]. For pulse addi-
`tion it has been recommended that 80% of the maximum
`refolding yield should be reached before adding the next
`pulse [14]. Other factors to be considered are the increas-
`
`ing residual concentration of the denaturant with each
`pulse, which should not surpass concentrations that affect
`the refolding of the protein, and the amount of protein
`added per pulse, which should be optimized in batch
`experiments to minimize aggregation [14].
`
`Membrane controlled denaturant removal
`Another technique to transfer the solubilized and
`unfolded protein to conditions allowing the formation of
`the native structure is the utilization of dialysis and diafil-
`tration systems for denaturant removal [e.g. [48-51]]. In
`contrast to the direct dilution method, the change from
`denaturing to native buffer conditions occurs gradually.
`Thus, the protein passes through different regimes of
`denaturant concentrations, where folding intermediates
`that are prone to aggregation may become populated.
`Most often, these techniques cause more aggregation dur-
`ing refolding compared to the direct dilution method [e.g.
`[52]]. Additionally, refolding yields can be negatively
`affected by non-specific adsorption of protein to the
`membrane. However, for some proteins and with the
`appropriate denaturant removal rates, adapted to the
`requirements of the target protein, high refolding yields at
`high protein concentrations can be obtained [50-53]. A
`fairly simple device was recently introduced allowing con-
`tinuous or pulse refolding in a similar way as in the direct
`dilution method [54].
`
`Chromatographic methods for protein refolding
`Protein refolding based on size exclusion chromatography
`Buffer exchange for denaturant removal can also be car-
`ried out by using size exclusion chromatography (SEC).
`Most frequently, the denaturant-protein solution is
`injected into a column previously equilibrated with the
`refolding buffer [44,55-58]. Subsequent elution with the
`refolding buffer results in a refolded protein in the eluate
`fraction with a considerably higher concentration com-
`pared to concentrations that can be reached by the simple
`dilution technique [44,56,58]. Protein refolding may be
`completed in the column or for proteins with slow folding
`kinetics the final folding steps may occur in the eluate
`fraction [44]. Aggregate formation is supposed to be
`reduced either by physical separation of aggregation-
`prone folding intermediates in the porous structures of
`the gel [56] or, more likely, by resolubilization of formed
`aggregates through the delayed running front of the
`denaturant, which gives the solubilized aggregates
`another opportunity to refold [14]. For proteins, which
`exhibit superior refolding yields during gradual denatu-
`rant removal, such as lysozyme, elution during SEC is
`preferably performed by using a decreasing denaturant
`gradient [38,52,59]. In specific cases, the denaturant
`removal can be accompanied with other changes in the
`buffer composition (i.e. pH) for further optimization of
`refolding conditions [52]. An additional advantage of this
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`chromatographic method is the concomitant purification
`of the target protein during the refolding process [44].
`Furthermore, some recent applications have shown the
`feasibility of using SEC for continuous processes of pro-
`tein refolding [60,61]. Also, SEC in combination with the
`use of an annular chromatography system can be coupled
`to an ultrafiltration and recycling unit for reinjection of
`resolubilized aggregates, which may form during the
`refolding process [60].
`
`Some parameters for refolding using SEC are of key
`importance. For example, protein aggregation during
`sample injection can cause low refolding yields [62];
`injecting the sample followed by an additional small vol-
`ume of denaturant solution solves
`this problem
`[44,52,62]. Also, optimum results can only be reached
`when the properties of the chromatographic resin allow
`efficient separation of the renatured target protein from
`different folding intermediates, misfolded protein, and
`aggregates that might form during the refolding process
`[59,63,64]. In general, lower refolding yields are obtained
`by injecting the denatured protein at high concentrations
`[38,58,60,61,64] and/or by elution at high rates
`[52,62,64]. Both conditions result in poor separation
`among different folding intermediates thereby boosting
`protein precipitation.
`
`Matrix-assisted protein refolding
`Attaching the solubilized and unfolded protein to a solid
`support prior to changing from denaturing to native
`buffer conditions is another approach to avoid the
`unwanted intermolecular interaction between aggrega-
`tion-prone folding intermediates. Binding of the solubi-
`lized and unfolded protein to the matrix requires the
`formation of a stable protein-matrix complex withstand-
`ing the presence of chaotropic agents. However, after
`changing to native buffer conditions, the detachment of
`the refolded target protein from the matrix should easily
`be accomplished. Several combinations of binding
`motives and matrices have been employed for binding the
`unfolded protein to the solid support. For example, pro-
`teins with a natural occurring charged patch in the
`unfolded chain, which binds to ion exchange resins
`[59,65-67], or proteins containing artificially engineered
`peptide tags such as the hexahistidine tag, which binds to
`immobilized metal ions [59,68-70], or N- or C-terminal
`hexaarginine tags binding to a polyanionic support [71],
`or protein fusions with denaturant-resistant binding
`domains, such as a glutathione S-transferase fragment,
`which binds to an anion exchange matrix [72] or the cel-
`lulose binding domain of the cellulose degrading mul-
`tienzyme complex of
`the
`thermophilic bacterium
`Clostridium thermocellum, which binds to a cellulose
`matrix [73], have been employed. After binding, the
`matrix-protein complex is brought to refolding conditions
`
`by any of the above-mentioned techniques such as dilu-
`tion [71], dialysis [68,73], or buffer exchange through
`chromatography [59,66,69,70,72]. Finally, the refolded
`protein can be detached from the matrix, e.g. in the case
`hexahistidine-tagged proteins by elution with EDTA [69]
`or imidazole [59,70] or by buffers with high ionic
`strength in the case of proteins bound by ionic interac-
`tions [59,65,66,71,72]. Due to the selective binding,
`matrix-assisted refolding can combine the renaturation of
`the target protein along with its purification from host cell
`protein contaminants [69,70,72].
`
`Refolding using hydrophobic interaction chromatography
`Hydrophobic interaction chromatography (HIC) has also
`been successfully used for protein refolding with concom-
`itant removal of contaminating proteins during the rena-
`turation process [74-78]. Unfolded proteins are applied to
`the column at high salt concentrations and refolded and
`eluted with a decreasing salt gradient. In contrast to the
`above-mentioned chromatographic methods there is no
`requirement for typical refolding aiding agents such as
`arginine during the in-column refolding process. Moreo-
`ver, refolding of the disulfide containing protein proinsu-
`lin was even obtained in the absence of a redox system in
`the mobile phase [76].
`
`It has been proposed that refolding is facilitated during
`HIC because unfolded proteins adsorb at high salt con-
`centrations to the hydrophobic matrix and, thus, are not
`prone to aggregation. Additionally, hydrophobic regions
`of the protein that adsorb to the HIC matrix form micro-
`domains around which native structure elements can
`form. During migration through the column, the protein
`will pass through several steps of adsorption and desorp-
`tion, controlled by the salt concentration and hydropho-
`bicity of the intermediate(s), resulting finally in the
`formation of the native structure [75].
`
`Physical and chemical features improving protein refolding
`yields
`Apart from any of the above-mentioned techniques for
`protein refolding, there are physical and chemical varia-
`bles that have a great impact on the final yield of biologi-
`cally active protein. For example, temperature as well as
`the composition of the refolding buffer are important var-
`iables influencing the final refolding yield.
`
`Physical variables aiding protein refolding
`The most important physical variable influencing the
`refolding yield is the temperature [40,45,50,51]. Temper-
`ature has a dual effect on the refolding process. On one
`side, it influences the speed of folding and on the other it
`influences the propensity towards aggregation of folding
`intermediates with exposed hydrophobic patches. Also,
`there is limited temperature range in which each protein
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`is thermodynamically stable in a given buffer system [79].
`In general, low temperatures support the productive fold-
`ing pathway as hydrophobic aggregation is suppressed.
`However, low temperatures also slow down the folding
`rates, thus increasing the time required for renaturation
`[51]. For refolding attempts of a new protein, 15°C has
`been proposed as a good starting point [14].
`
`Pressure was identified as another important physical var-
`iable affecting protein structure as well as protein refold-
`ing processes [80]. It was shown that high pressure up to
`3 kbar can disrupt oligomeric protein structures [80] and
`can dissolve protein aggregates and inclusion bodies
`[81,82]. The disassembled protein monomers retain
`native-like secondary structure up to 5 kbar [80]. After
`gradual depressurization, they can reach their native state
`even at high protein concentrations, because folding
`intermediates prone to aggregate at atmospheric pressure
`are prevented from aggregation by high pressure [81-83].
`
`Chemicals aiding protein refolding
`Certainly, L-arginine is nowadays the most commonly
`used refolding aiding agent [14]. It impedes aggregate for-
`mation by enhancing the solubility of folding intermedi-
`ates, presumably by shielding hydrophobic regions of
`partially folded chains. In addition, it has been shown
`that numerous other low molecular weight additives such
`as detergents, protein-stabilizing agents such as glycerol or
`even low residual concentrations of denaturants improve
`refolding yields by suppressing aggregation [14]. In addi-
`tion, high-molecular weight additives such as polyethyl-
`ene glycol were used successfully for enhancing protein
`refolding yields [84]. More recently, low-molecular
`weight non-detergent zwitterionic agents such as sulfo-
`betaines, substituted pyridines and pyrroles and acid sub-
`stituted aminocyclohexanes have been employed
`successfully for protein renaturation [40,85-87]. Moreo-
`ver, polymers with temperature-dependent hydrophobic-
`ity were effectively applied for protein refolding at higher
`temperatures [88,89]. The benefit of each of these refold-
`ing aiding agents for a given renaturation system has to be
`elucidated experimentally, as they are not equally advan-
`tageous for all proteins. The mechanisms of interactions
`of these refolding aiding agents with the folding interme-
`diates remain often obscure although it is clear that all
`these substances suppress aggregation in favor of the pro-
`ductive folding pathway [90].
`
`Micelles and liposomes as protein refolding aiding systems
`Detergents [91,92] and phospholipids [93,94], in the
`form of micelles and liposomes [95], respectively, as well
`as mixed micelle systems formed by phospholipids and
`detergents [92,95,96] have shown potential to aid protein
`refolding. Most likely, illegitimate hydrophobic interac-
`tions between folding intermediates are suppressed by
`
`transient nonpolar interactions between the protein and
`the micelle or liposome [91-93]. Additional transient
`polar interactions in mixed micelles are supposed to be
`responsible for higher refolding yields compared to only
`detergent-based micelle systems [92,96]. Moreover, lipo-
`somes linked covalently to chromatographic resins have
`potential to combine renaturation and separation of the
`refolded target protein [93,94].
`
`Reversed micelles, formed when an aqueous detergent
`solution is mixed with an organic solvent, can also facili-
`tate protein refolding by avoiding aggregate formation
`[97]. The denatured protein, once transferred to this solu-
`tion, tries to avoid the organic phase, and, after reaching
`the hydrophilic core of the reversed micelle, can refold as
`a single molecule [97]. Recently, it was demonstrated that
`protein precipitates can be solubilized by direct addition
`into the reversed micellar system allowing refolding with
`high yields at high protein concentrations [98-100]. Yet,
`direct solubilization of inclusion bodies in reversed micel-
`lar systems has not been reported. In addition, recovery of
`refolded protein from these micellar structures is not eas-
`ily accomplished [97,99].
`
`Chemical and biological protein refolding aiding agents
`mimicking in vivo folding conditions
`Natural chaperones
`Chaperones are a group of proteins conserved in all king-
`doms, which play a key role in assisting in vivo protein
`folding and protecting cellular proteins from different
`types of environmental stress by suppressing protein
`aggregation. For example, the major E. coli chaperonin
`GroEL is involved in the in vivo folding of 10% of all newly
`synthesized proteins at normal growing conditions, and
`of 30% under stress conditions [101]. GroEL assists pro-
`tein folding by a first capturing step of aggregation-prone
`folding intermediates [102]. The release of the folding-
`competent form is then accomplished in an ATP-depend-
`ent fashion through the action of the cochaperonin GroES
`[102].
`
`Natural chaperones have also been applied successfully to
`refold various proteins in vitro [103]. However, their rou-
`tine application is limited by their cost, the relatively high
`chaperone concentration required (at least equimolar to
`the target protein) and the need for their removal after the
`refolding procedure [103,104]. Some procedures have
`tried to overcome these limitations by utilizing immobi-
`lized and reusable (mini)-chaperone systems [104-106].
`Nevertheless, chaperone-based refolding processes are not
`robust enough for large-scale processes [14].
`
`Artificial chaperones
`A further development of the detergent-based micellar
`system mimics the two-step mechanism of chaperone-
`
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`Microbial Cell Factories 2004, 3:11
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`http://www.microbialcellfactories.com/content/3/1/11
`
`assisted protein folding. The capturing step is performed
`by diluting the denatured protein into a detergent solu-
`tion, which prevents protein aggregation through the for-
`mation of mixed protein-detergent micelles [107-110].
`Aqueous solutions of hydrogel nanoparticles (e.g. self-
`assembly of hydrophobized polysaccharides such as cho-
`lesterol-bearing pullulan) have been also used for the cap-
`turing step [111]. The release of the folding-competent
`protein is subsequently initiated by the addition of cyclo-
`dextrins [107-112]. They are added in excess to the