202
`
`Protein refolding for industrial processes
`Eliana De Bernardez Clark
`
`Inclusion body refolding processes are poised to play a major
`role in the production of recombinant proteins. Improving
`renaturation yields by minimizing aggregation and reducing
`chemical costs are key to the industrial implementation of
`these processes. Recent developments include solubilization
`methods that do not rely on high denaturant concentrations
`and the use of high hydrostatic pressure for simultaneous
`solubilization and renaturation.
`
`Addresses
`Department of Chemical and Biological Engineering, Tufts University,
`Medford, MA 02155, USA; e-mail: eliana.clark@tufts.edu
`
`Current Opinion in Biotechnology 2001, 12:202–207
`
`0958-1669/01/$ — see front matter
`© 2001 Elsevier Science Ltd. All rights reserved.
`
`Abbreviations
`CTAB
`n-cetyl trimethylammonium bromide
`DTE
`dithioerythritol
`DTT
`dithiothreitol
`GdmCl guanidinium chloride
`PDGF
`platelet-derived growth factor
`SDS
`sodium dodecyl sulfate
`SEC
`size-exclusion chromatography
`
`Introduction
`The need for the efficient production of genetically engi-
`neered proteins has grown and will continue to grow as a
`consequence of the success of the human genome project. A
`variety of hosts may be used to produce these proteins, with
`expression in bacteria poised to play a major role, particular-
`ly when the biological activity of the protein product is not
`dependent on post-translational modifications. Expression
`of genetically engineered proteins in bacteria often results
`in the accumulation of the protein product in inactive insol-
`uble deposits inside the cells, called inclusion bodies. Faced
`with the prospect of producing an insoluble and inactive
`protein, researchers usually attempt to improve solubility by
`a variety of means, such as growing the cells at lower tem-
`peratures, co-expressing the protein of interest with
`chaperones and foldases and using solubilizing fusion part-
`ners, among others [1]. However, expressing a protein in
`inclusion body form can be advantageous. Large amounts of
`highly enriched proteins can be expressed as inclusion bod-
`ies. Trapped in insoluble aggregates, these proteins are for
`the most part protected from proteolytic degradation. If the
`protein of interest is toxic or lethal to the host cell, then
`inclusion body expression may be the best available pro-
`duction method. The challenge is to take advantage of the
`high expression levels of inclusion body proteins by being
`able to convert inactive and misfolded inclusion body
`proteins into soluble bioactive products [2–5].
`
`The recent literature includes many examples of the
`refolding of genetically engineered proteins. A significant
`
`number of these publications deal with the expression
`and purification of small amounts of proteins for structure/
`function relationship and biophysical characterization
`studies. Although valuable, the processes described in
`these publications are usually inefficient, include multiple
`unnecessary steps and have very low recovery yields. A
`second significant fraction of the refolding literature deals
`with understanding the folding pathway of a variety of pro-
`teins and, in particular, early folding events. These studies
`are performed with purified proteins that are subjected to
`unfolding under a variety of conditions, followed by
`carefully designed and monitored refolding experiments.
`A third fraction of the refolding literature, and the focus of
`this review, deals with the development of more efficient
`refolding methods that can be used for the commercial
`production of genetically engineered proteins
`
`The general strategy used to recover active protein from
`inclusion bodies involves three steps: inclusion body isola-
`tion and washing; solubilization of the aggregated protein;
`and refolding of the solubilized protein (Figure 1a).
`Although the efficiency of the first two steps can be rela-
`tively high, renaturation yields may be limited by the
`accumulation of inactive misfolded species as well as aggre-
`gates. Because the majority of industrially relevant proteins
`contain one or more disulfide bonds, this review focuses on
`recent advances in oxidative protein refolding, that is,
`refolding with concomitant disulfide-bond formation.
`
`Inclusion body isolation, purification and
`solubilization
`Inclusion bodies are dense, amorphous protein deposits that
`can be found in both the cytoplasmic and periplasmic space
`of bacteria [1,6•]. Structural characterization studies using
`ATR-FTIR (attenuated total reflectance Fourier-trans-
`formed infrared spectroscopy) have shown that the
`insoluble nature of inclusion bodies may be due to their
`increased levels of non-native intermolecular β-sheet con-
`tent compared with native and salt-precipitated protein
`[7,8]. Cells containing inclusion bodies are usually disrupted
`by high-pressure homogenization or a combination of
`mechanical, chemical and enzymatic methods [6•,9•]. The
`resulting suspension is treated by either low-speed centrifu-
`gation or filtration to remove soluble proteins from the
`particulate containing the inclusion bodies. The most
`difficult to remove contaminants of inclusion body protein
`preparations are membrane-associated proteins that are
`released upon cell breakage. Washing steps are performed to
`remove membrane proteins and other contaminants.
`Methods used to solubilize prokaryotic membrane proteins
`can be adapted to wash inclusion bodies. The most common
`washing steps utilize EDTA, and low concentrations of
`denaturants and/or weak detergents such as Triton X-100,
`deoxycholate and octylglucoside [6•,9•,10,11•,12,13,P1,P2].
`
`Page 1
`
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`

`Batas, Schiraldi and Chaudhuri [10] recently compared
`centrifugation and membrane filtration for the recovery
`and washing of inclusion bodies. Two membrane pore sizes
`(0.1 and 0.45 µm) were compared; the larger pore size
`membrane gave better solvent flux and protein purity.
`Centrifugation resulted in higher protein purity, probably
`because it takes advantage of the density differences
`between cell debris and inclusion bodies.
`
`A variety of methods may be used to solubilize inclusion
`bodies; however, the choice of solubilizing agent can great-
`ly impact the subsequent refolding step and the cost of the
`overall process. The most commonly used solubilizing
`agents are denaturants, such as guanidinium chloride
`(GdmCl) and urea. Using these denaturants, solubilization
`may be accomplished by the complete disruption of the
`protein structure (unfolding) or by the disruption of inter-
`molecular interactions with partial unfolding of the
`protein. The latter approach has the advantage that it
`requires lower amounts of denaturant to succeed.
`Although proteins have been successfully refolded from
`the denatured state, it may prove to be difficult to fold pro-
`teins from a partially folded state. Key to the development
`of an efficient and economic denaturant-based solubiliza-
`tion step is the determination of the minimum amount of
`denaturant needed to solubilize the protein and to allow
`for full bioactivity recovery in the refolding step. The
`majority of the published work on inclusion body protein
`refolding has used relatively high denaturant (6–8 M) and
`protein (1–10 mg/ml) concentrations in the solubilization
`step [5,9•,10,11•,12–14].
`
`Lower denaturant concentrations (1–2 M) have been used
`to solubilize cytokines from Escherichia coli inclusion bod-
`ies [P3]. The purity of the solubilized protein was much
`higher at GdmCl concentrations of 1.5–2 M compared with
`the more commonly used 4–6 M concentrations, because
`at the higher GdmCl concentrations contaminating pro-
`teins were also released from the particulate fraction. No
`information was provided about the efficiency of this solu-
`bilization process or the range of inclusion body protein
`concentrations for successful solubilization.
`
`Extremes of pH have also been used to solubilize inclusion
`bodies. Gavit and Better [15] used a combination of low pH
`(≤ 2.6) and high temperature (85°C) to solubilize antifungal
`recombinant peptides from E. coli. Lower temperatures and
`higher pH values resulted in increased solubilization time.
`Reddy and coworkers [16] utilized 20% acetic acid to solu-
`bilize a maltose-binding protein fusion from inclusion
`bodies. These low pH solubilization processes may not be
`applicable to many proteins, particularly those that undergo
`irreversible chemical modifications at these conditions or
`those susceptible to acid cleavage.
`
`High pH (≥ 12) has been used to solubilize growth hormones
`[17,18] and proinsulin [P4]. Exposure to elevated pH condi-
`tions for extended periods of time may also cause irreversible
`
`Protein refolding for industrial processes De Bernardez Clark 203
`
`Figure 1
`
`(a) Cells containing inclusion bodies
`
`Homogenization
`Centrifugation or microfiltration
`
`Soluble fraction
`
`Particulate
`Washing
`Solubilization
`Centrifugation or microfiltration
`
`Soluble proteins
`
`Debris
`
`Renaturation with or without
`prior purification
`
`Active protein
`
`(b) Cells containing inclusion bodies
`
`In situ solubilization
`Centrifugation or microfiltration
`
`Soluble fraction
`
`Debris
`
`Purification
`Renaturation
`
`Active protein
`
`Current Opinion in Biotechnology
`
`Processes for the recovery of inclusion body proteins. (a) Inclusion
`body isolation followed by solubilization. (b) The in situ solubilization of
`inclusion bodies.
`
`chemical modifications to the protein. Thus, this high pH
`solubilization method, although attractive for its simplicity
`and low cost, may not be applicable to most pharmaceutical
`proteins. More effective solubilization methods for growth
`hormones combine high pH with low denaturant concentra-
`tions [17,18], 20–40% isopropyl or n-propyl alcohol solutions
`[P1] or acyl glutamate detergents [P5].
`
`Detergents have also been used to solubilize inclusion bodies.
`Commonly used detergents are sodium dodecyl sulfate
`(SDS) and n-cetyl trimethylammonium bromide (CTAB)
`[3,18,19]. Detergents offer the advantage that the solubilized
`protein may already display biological activity, thus avoiding
`the need for a refolding step. If this is the case, it is important
`to remove contaminating membrane-associated proteases in
`the inclusion body washing step to avoid proteolytic degrada-
`tion of the solubilized inclusion body protein [6•]. One
`acknowledged drawback of the use of detergents as solubiliz-
`ing agents is that they may interfere with downstream
`chromatographic steps. Extensive washing may be needed to
`remove solubilizing detergents [P5]. Alternatively, detergents
`may be extracted from refolding mixtures by using cyclodex-
`trins [20], linear dextrins [21] or cycloamylose [22].
`
`Page 2
`
`

`

`204 Biochemical engineering
`
`Patra and coworkers [18] compared several solubilization
`methods for the recovery of human growth hormone from
`E. coli inclusion bodies. They observed similar solubiliza-
`tion efficiencies when using 8 M urea, 6 M GdmCl, 1%
`SDS or 1% CTAB (all at pH 8.5) or 2 M urea (at pH 12.5).
`Refolding for the first four solubilization conditions
`required a dilution step resulting in increased process vol-
`umes. Solubilization in 2 M urea at pH 12.5 was simple,
`economical and efficient, and refolding could be accom-
`plished by a simple pH adjustment without dilution.
`However, this high pH solubilization method may not be
`applicable to proteins that might undergo irreversible
`chemical modifications under these conditions.
`
`A key to the solubilization process is the addition of a
`reducing agent to maintain cysteine residues in the
`reduced state and thus prevent non-native intra- and inter-
`disulfide bond formation in highly concentrated protein
`solutions at alkaline pH. Typically used reducing agents
`are dithiothreitol (DTT), dithioerythritol (DTE), and
`2-mercaptoethanol [2,3]. These reducing agents should be
`added in slight excess to ensure complete reduction of all
`cysteine residues. Chelating agents are added to the solu-
`bilization solution to prevent metal-catalyzed air oxidation
`of cysteines. Alternatively, reduced cysteines may be pro-
`tected from oxidation by the formation of S-sulfonate
`derivatives [23,P6,P7] or mixed disulfides [9•,P7].
`
`When expression levels are very high, a competitive alter-
`native is to add the solubilizing agents directly to the broth
`at the end of the fermentation process. This in situ solubi-
`lization method has been used to recover insulin-like
`growth factor using urea under alkaline conditions [P8] and
`antifungal recombinant peptides using a combination of
`low pH (< 2.6) and high temperature (85°C) [15]. The
`main disadvantage of in situ solubilization concerns the
`release of both proteinaceous and nonproteinaceous conta-
`minants that may have to be removed before renaturation
`is attempted. It has been shown that protein refolding in
`the presence of impurities may result in decreased yields
`[6•,24]. The main advantage of this method is the elimina-
`tion of time-consuming and energy-consuming mechanical
`disruption methods and of one centrifugation and/or
`filtration step (Figure 1b).
`
`Solubilization may also be accomplished by applying high
`hydrostatic pressures (1–2 kbar) in the presence of reducing
`agents and low concentrations of solubilizing agents [25•,P9].
`
`Renaturation of the solubilized protein
`When inclusion bodies have been solubilized using a com-
`bination of reducing agents and high concentrations of
`denaturants, renaturation is then accomplished by the
`removal of excess denaturants by either dilution or a
`buffer-exchange step, such as dialysis, diafiltration, gel-fil-
`tration chromatography or immobilization onto a solid
`support. Because of its simplicity, dilution of the solubi-
`lized protein directly into renaturation buffer is the most
`
`commonly used method in small-scale refolding studies.
`The main disadvantages of dilution refolding for commer-
`cial applications are the need for larger refolding vessels
`and additional concentration steps after renaturation. The
`key to successful dilution refolding is to control the rate of
`the addition of denatured protein to renaturation buffer
`and to provide good mixing in order to maintain low pro-
`tein concentration during refolding and thus prevent
`aggregation. Dilution refolding can also be accomplished
`in multiple steps, also known as ‘pulse renaturation’, in
`which aliquots of denatured reduced protein are added to
`renaturation buffer at successive time intervals [2,9•], or
`semicontinuously via fed-batch addition of the denatured
`reduced protein to refolding buffer [26]. Recently, Katoh
`and Katoh [26] developed a continuous refolding method
`in which denatured reduced protein is gradually added
`from the annular space of a membrane tube to renaturation
`buffer flowing continuously through the inner space of the
`membrane tube. Refolding yields obtained using this con-
`tinuous refolding method were similar to those obtained
`using fed-batch dilution and about 10% higher than those
`using batch dilution [26].
`
`Buffer exchange to remove high denaturant concentrations
`can also be accomplished by diafiltration [27] and dialysis
`[28] using ultrafiltration membranes. Renaturation yields
`using these membrane-based methods can be significantly
`affected by protein binding to the membranes. Binding
`can be minimized by using highly hydrophilic materials,
`such as cellulose acetate, which are more compatible with
`unfolded protein molecules. With typical hydrophobic
`membrane materials, such as polyether sulfone, the major-
`ity of the denatured protein was found bound to the
`membrane [28]. Significant losses of unfolded protein
`occurred via transmission through the membrane. These
`losses could be reduced by dialysis against lower denatu-
`rant concentrations that lead to molten-globule or native
`configurations [28].
`
`Size-exclusion chromatography (SEC) is an alternative
`buffer-exchange method to remove high denaturant con-
`centrations and promote renaturation [11•,13,29•]. Fahey,
`Chaudhuri and Binding [13] examined the effect of gel
`type on renaturation yields and found that as the fraction-
`ation range of the gel matrix increases from Sephacryl
`(S)-100 to S-400, aggregation decreases but the resolution
`between protein and denaturant decreases. Thus, opti-
`mum renaturation yields were obtained with the S-300 gel.
`In a separate study, Fahey, Chaudhuri and Binding [29•]
`compared batch dilution and SEC refolding. Dilution also
`takes place during SEC but, for similar dilution factors,
`SEC resulted in higher refolding yields when compared
`with batch dilution as long as the dilution factor was below
`40. Sample application conditions were found to have a
`strong effect on the efficiency of SEC refolding, because
`rapid structural collapse takes place during sample applica-
`tion that can lead to aggregation. Renaturation yields
`decreased with higher protein concentrations and sample
`
`Page 3
`
`

`

`volumes and lower flow rates [29•]. Muller and Rinas [11•]
`circumvented the problem of aggregation during sample
`application by allowing the denatured protein to penetrate
`the column under denaturing conditions and then chang-
`ing
`the buffer
`to
`renaturation conditions. They
`successfully refolded the complex heterodimeric protein
`platelet-derived growth factor (PDGF) using a combina-
`tion of SEC for refolding of the monomeric species
`followed by prolonged incubation under renaturation
`conditions to promote dimerization.
`
`Buffer exchange to remove high denaturant concentrations
`can also be achieved by transiently binding the denatured
`protein to a solid support. Intermolecular interactions lead-
`ing to aggregation are minimized when the refolding
`molecules are isolated through binding to the support.
`Freedom for structure formation during renaturation is
`facilitated by binding through fusion partners, such as a
`His-tag [30] or the cellulose-binding domain [31], which
`retain their binding capabilities under the denaturing con-
`ditions required for loading the solubilized inclusion body
`protein onto the column. In situ purification is achieved by
`washing the bound protein before elution.
`
`Disulfide-bond formation during folding
`In the case of disulfide-bonded proteins, renaturation
`buffers must promote disulfide-bond formation (oxidation).
`The simplest and most inexpensive oxidation method uses
`air in the presence of a metal catalyst and a reducing agent
`to facilitate disulfide-bond reshuffling [P1,P8]. The rate of
`disulfide-bond formation through air oxidation may be lim-
`ited by the slow mass transfer rate of oxygen in aqueous
`solutions. Increased agitation, which can be used to improve
`mass transfer rates, may also lead to aggregation due to
`increased shear and interfacial stresses [32].
`
`Oxidation rates can be accelerated using an oxido-shuffling
`system, which consists of mixtures of reduced and oxidized
`low molecular weight thiol reagents. The most commonly
`used oxido-shuffling reagents are reduced and oxidized
`glutathione (GSH/GSSG) but the pairs cysteine/cystine,
`cysteamine/cystamine, DTT/GSSG and DTE/GSSG have
`also been utilized [2,3,9•]. Molar ratios of reduced to oxi-
`dized thiol of 3:1 to 1:1 and total thiol concentrations
`between 5–15 mM have been found to be optimal [14,33].
`Disulfide-bond formation using the oxido-shuffling system
`can be accelerated by using a small-molecule mimic of pro-
`tein disulfide isomerase [34,P10]. A disadvantage of the
`oxido-shuffling system is the high cost of some of the
`reagents, particularly glutathione.
`
`A third oxidation method uses a two-step approach: the
`formation of mixed disulfides between glutathione and the
`denatured protein before renaturation, followed by refold-
`ing in the presence of catalytic amounts of a reducing
`agent to promote disulfide-bond formation and reshuffling
`[9•,P7]. The refolding yield of recombinant human tissue
`plasminogen activator could be increased sixfold when the
`
`Protein refolding for industrial processes De Bernardez Clark 205
`
`oxido-shuffling refolding system was replaced with the
`mixed disulfide approach [9•]. Alternatively, cysteines in
`the denatured protein may be protected by sulfonation,
`followed by the addition of a reducing agent such as
`cysteine [P7] and 2-mercaptoethanol [P6] or a thiol/
`disulfide mixture such as cysteamine/cystamine [23].
`
`Improving renaturation yields
`The formation of incorrectly folded species, and in partic-
`ular aggregates, is usually the cause of decreased
`renaturation yields. A very efficient strategy to suppress
`aggregation is the inhibition of the intermolecular interac-
`tions leading to aggregation by the use of low molecular
`weight additives. These small molecules are relatively
`easy to remove when refolding is complete. Numerous
`additives have been shown to prevent aggregation [3,9•].
`The mechanism of action of additives is still unclear. They
`may influence both the solubility and the stability of the
`native, denatured and intermediate state(s), they may act
`by changing the ratio of the rates of proper folding and
`aggregate formation or they might simply act by solubiliz-
`ing aggregates. The most commonly used low molecular
`weight additives are L-arginine (0.4–1 M), low concentra-
`tions of denaturants such as urea (1–2 M) and GdmCl
`(0.5–1.5 M) and detergents (Chaps, SDS, CTAB and
`Triton X-100). In a recent review, De Bernardez Clark,
`Schwarz and Rudolph [9•] discussed different approaches
`to inhibiting aggregation during refolding and provided a
`detailed list of low molecular weight additives and the con-
`centration ranges needed to increase renaturation yields.
`
`Low concentrations of urea and GdmCl, although widely
`used to inhibit aggregation [14,29•,P2,P3,P7], are not
`always effective folding enhancers. GdmCl concentrations
`as low as 0.25 M were found to inhibit the oxidative dimer-
`ization of PDGF [11•]. Similarly, bone morphogenetic
`protein-2 proved difficult to refold in the presence of low
`concentrations of denaturants [35].
`
`Detergents such as Chaps [12,35], CTAB [20,21], Triton
`X-100 [20] and SDS [19] have been successfully used to
`improve renaturation yields. As noted earlier under solubi-
`lization methods, one drawback of the use of detergents is
`that they may be difficult to remove and may affect down-
`stream chromatographic steps. Detergents have been
`extracted from refolding mixtures using cyclodextrins [20],
`linear dextrins [21], cycloamylose [22] and ion-exchange
`chromatography in the case of ionic detergents [19].
`
`The in vivo competition between folding and aggregation
`is modulated by chaperones and foldases [1]. It is not
`surprising that these proteins can also affect the in vitro
`competition between folding and aggregation [36].
`Because chaperones and foldases are proteins that need to
`be removed from the renaturation solution at the end of
`the refolding process and as they may be costly to pro-
`duce, their commercial use will require a recovery–reuse
`scheme. Altamirano and coworkers [37] developed a
`
`Page 4
`
`

`

`206 Biochemical engineering
`
`Table 1
`
`Most frequently used oxidative renaturation methods.
`
`Solubilization method
`
`Renaturation method*
`
`Comments
`
`High denaturant concentrations:
`urea (≥ 8 M) or GdmCl
`(≥ 6 M) in the presence of a reducing
`agent (DTT, DTE, 2-mercaptoethanol)
`
`Detergents:
`SDS, CTAB
`in the presence of a reducing agent
`(DTT, DTE, 2-mercaptoethanol)
`
`Removal of the denaturant by dilution or
`buffer exchange (dialysis, diafiltration,
`size-exclusion chromatography or binding
`to a matrix) with concomitant disulfide-bond
`formation†
`
`Oxidation† followed by detergent removal
`by extensive washing or stripping
`
`Residual denaturant concentrations
`may interfere with the assembly of
`oligomeric proteins
`
`References
`[9•,11•,12–14,
`18,29•,P2]
`
`Residual detergent concentrations
`may interfere with downstream
`purification processes
`
`[18–22]
`
`Extremes of pH in the presence of
`low concentrations of denaturants
`
`pH adjustment to 7.5–9.5 to promote
`disulfide-bond formation†
`
`Extreme pH may cause irreversible
`chemical modifications
`
`[15–18]
`
`*Low molecular weight additives may be added to prevent aggregation. †Disulfide-bond formation may be accomplished by air oxidation in the
`presence of a metal catalyst and a reducing agent or by using an oxido-shuffling system.
`
`reusable molecular chaperone system for oxidative refold-
`ing chromatography that utilizes a GroEL minichaperone,
`which can prevent aggregation, the oxido-shuffling cata-
`lyst DsbA, and peptidyl-prolyl isomerase, all immobilized
`on an agarose gel. Recently, Kohler, Preuss and Miller [38]
`developed a chaperone-assisted refolding bioreactor that
`uses a stirred-cell membrane system to immobilize the
`GroEL–GroES complex. In its current design, the biore-
`actor could only be used for three cycles of refolding.
`Further design improvements will be needed before this
`bioreactor can be considered as a commercially viable
`refolding alternative.
`
`Interestingly, high hydrostatic pressures (1–2 kbar) in
`combination with low concentrations of denaturants
`have been used for the simultaneous solubilization and
`refolding of inclusion body proteins [25•,P9]. Similarly,
`high hydrostatic pressures can be used in the refolding
`process to prevent aggregation [39,40].
`
`Conclusions
`The recovery of bioactive proteins from inclusion bodies
`is a complex process. Despite its complexity, there are
`clear guidelines on how to proceed when faced with the
`task of refolding an inclusion body protein (Figure 1;
`Table 1). As with other protein recovery processes, how-
`ever, optimum conditions have to be determined on a
`case by case basis. The key to a commercially viable
`renaturation process lies in minimizing the number of
`steps (to increase the overall yield) and the amounts and
`costs of chemicals needed. This can be accomplished by
`eliminating unnecessary buffer-exchange steps, by
`exploring the use of alternative solubilization methods
`that do not rely on high denaturant concentrations, and
`by developing efficient oxidation methods that do not
`require the use of expensive oxido-shuffling systems.
`Future developments in protein refolding will benefit
`from a more fundamental understanding of inclusion
`body solubilization methods, and on the role that addi-
`tives play in the inhibition of aggregation.
`
`1.
`
`2.
`
`3.
`
`4.
`
`References and recommended reading
`Papers of particular interest, published within the annual period of review,
`have been highlighted as:
`(cid:127) of special interest
`(cid:127)(cid:127) of outstanding interest
`Baneyx F: Recombinant protein expression in Escherichia coli.
`Curr Opin Biotechnol 1999, 10:411-421.
`Lilie H, Schwarz E, Rudolph R: Advances in refolding of proteins
`produced in E. coli. Curr Opin Biotechnol 1998, 9:497-501.
`De Bernardez Clark E: Refolding of recombinant proteins. Curr
`Opin Biotechnol 1998, 9:157-163.
`Rudolph R, Lilie H, Schwarz E: In vitro folding of inclusion body
`proteins on an industrial scale. In Biotechnology, vol 5a. Eds
`Rehm H-J, Reed G. Weinheim, Germany: Wiley-VCH; 1999.
`5. Misawa S, Kumagai I: Refolding of therapeutic proteins produced
`in Escherichia coli as inclusion bodies. Biopolymers 1999,
`51:297-307.
`6. Georgiou G, Valax P: Isolating inclusion bodies from bacteria.
`•
`Methods Enzymol 1999, 309:48-58.
`This paper presents an excellent discussion of inclusion body formation and
`characterization. It includes methods and protocols for the isolation and
`purification of inclusion bodies from bacteria.
`Fink A: Protein aggregation: folding aggregates, inclusion bodies
`and amyloid. Fold Des 1998, 3:R9-R23.
`Seshadri S, Khurana R, Fink AL: Fourier transformed infrared
`spectroscopy in analysis of protein deposits. Methods Enzymol
`1999, 309:559-576.
`De Bernardez Clark E, Schwarz E, Rudolph R: Inhibition of
`aggregation side reactions during in vitro protein folding. Methods
`Enzymol 1999, 309:217-236.
`This article contains protocols for inclusion body isolation, purification and
`solubilization and for protein renaturation. It includes a detailed discussion of
`methods to inhibit aggregation during folding and a extensive list of low mol-
`ecular weight folding enhancers.
`10. Batas B, Schiraldi C, Chaudhuri JB: Inclusion body purification and
`protein refolding using microfiltration and size exclusion
`chromatography. J Biotechnol 1999, 68:149-158.
`11. Muller C, Rinas U: Renaturation of heterodimeric platelet-derived
`•
`growth factor from inclusion bodies of recombinant Escherichia
`coli using size-exclusion chromatography. J Chromatogr A 1999,
`855:203-213.
`This paper details a method for the successful expression and oxidative
`refolding of the complex heterodimeric protein platelet-derived growth factor
`using size-exclusion chromatography (SEC). The use of SEC allowed for the
`simultaneous refolding and purification of the protein. The effects of protein
`concentration, temperature, denaturant concentration, and thiol/disulfide
`concentrations during dimerization were examined. Maximum yields were
`obtained at 25°C, in the absence of denaturant and for a 10:1 ratio of
`
`7.
`
`8.
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`9.
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