Journal of Biotechnology 128 (2007) 587–596
`
`Review
`Current status of technical protein refolding
`∗
`
`Alois Jungbauer
`
`, Waltraud Kaar 1
`
`Department of Biotechnology, Austrian Center of Biopharmaceutical Technology, University of Natural Resources
`and Applied Life Sciences Vienna, Muthgasse 18, A-1190 Vienna, Austria
`
`Received 30 July 2006; received in revised form 14 November 2006; accepted 4 December 2006
`
`Abstract
`
`The expression of heterologous proteins in microbial hosts frequently leads to the formation of insoluble aggregates. To
`fully exploit the production capacity of the cells, efficient strategies for further processing have to be developed. While in lab
`scale matrix assisted refolding techniques, especially of histidine-tagged proteins have become very popular, in production scale
`refolding by dilution is still predominant due to its simplicity. However scaling up dilution processes leads to large volumes and
`low protein concentration. This is a heavy burden both for liquid handling and for subsequent downstream processing steps.
`Process development aims to operate at uniform, reproducible conditions, to reduce costs to a minimum and to guarantee the
`required quality of the product. The general refolding kinetics, exploration of appropriate refolding conditions are reviewed.
`The major refolding operations such as dilution, matrix assisted refolding, pressure driven refolding or continuous refolding
`applications are discussed in view of industrial applicability.
`© 2006 Elsevier B.V. All rights reserved.
`
`Keywords: Inclusion bodies; Escherichia coli; Refolding; Large scale
`
`Contents
`
`Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`1.
`2. Refolding kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`3.
`Isolation of recombinant protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`4. Determination of refolding conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`5. Refolding by dilution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`6.
`Pressure treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`
`588
`588
`590
`591
`592
`592
`
`∗
`
`Corresponding author. Fax: +43 1 3697615.
`E-mail address: alois.jungbauer@boku.ac.at (A. Jungbauer).
`1 Current address: Centre for Biomolecular Engineering, Australian Institute for Bioengineering and Nanotechnology, University of
`Queensland, St. Lucia 4072, Qld, Australia.
`
`0168-1656/$ – see front matter © 2006 Elsevier B.V. All rights reserved.
`doi:10.1016/j.jbiotec.2006.12.004
`
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`Large scale chromatographic refolding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`7.
`8. Analysis of folded proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`9. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
`
`593
`593
`594
`594
`
`1. Introduction
`
`Recombinant DNA technology allows the expres-
`sion of valuable heterologous proteins at high
`expression rates. Particularly in Escherichia coli (E.
`coli) overexpression of proteins often leads to aggrega-
`tion and deposition in dense, insoluble particles within
`the host cell, so-called inclusion bodies (IB). They
`are easily distinguishable from other cell components
`due to their refractile character (Fig. 1). Formation of
`inclusion bodies is heavily protein dependent, charge
`distribution and turn forming residues have a strong
`impact (Wilkinson and Harrison, 1991), also presence
`of cysteines may enforce tendency of aggregate forma-
`tion (Rinas et al., 1992) but it may also be influenced
`by altering cell cultivation conditions (Panda et al.,
`1999). Decelerated cell growth achieved by lower tem-
`perature (Schein and Noteborn, 1988) or suboptimal
`pH (Kopetzki et al., 1989) can result in the produc-
`tion of soluble recombinant proteins which indicates
`that the cells are overburdened by the protein produc-
`tion at regular cultivation conditions. Still production
`of proteins as inclusion bodies is favored in several
`cases. Intracellular expression of proteins does have
`
`certain advantages over secretion of the product into
`the culture supernatant. Design heuristics of biotech-
`nological processes recommend removing the most
`abundant impurities first. In a fermentation process
`this constitutes water. By a simple unit operation such
`as centrifugation the product can be concentrated by
`recovering the whole cells in the sediment. As outlined
`in Fig. 2 the attraction of inclusion body production
`compared to secretory systems is the simple primary
`recovery step. After cell harvest the cells have to be
`disintegrated and inclusion bodies have to be separated
`from cell debris and soluble cell components released
`into the homogenate.
`Inclusion bodies consist nearly exclusively of
`recombinant proteins (Speed et al., 1996). Isolation of
`the desired product at already high purity is relatively
`easy due to density differences (Schoner et al., 1985)
`and high protein concentration can be achieved at the
`primary solubilization step. Although there are studies,
`that inclusion body protein is not the dead end deposit
`as believed earlier but inclusion bodies are dynamic
`structures subjected to permanent conversion (Carrio
`and Villaverde, 2002), the storage as aggregates still
`features distinct protection from protease degradation
`(Cheng et al., 1981; Kitano et al., 1987). Another
`advantage is the possibility to produce compounds,
`which are cell toxic in higher concentration. However,
`as a major disadvantage, the subsequently required
`refolding procedure poses a bottleneck in every
`downstream scheme. Protein aggregates have to
`be resolved and folded into their native structure.
`Various strategies have been employed to achieve an
`active compound refolded from inclusion bodies in
`reasonable yield. In this review special emphasis is
`taken on the scalability of a method and the use in
`industrial production processes.
`
`2. Refolding kinetics
`
`Fig. 1. Electron micrograph of E. coli cells containing cytosolic
`inclusion bodies.
`
`The distinct folding pathway of a single protein is
`still case of many hypotheses. While debating the driv-
`
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`
`589
`
`Fig. 2. Generalized processing scheme for the production of protein formulation from fermentation broth. Bold lines depict a usual inclusion
`body processing route.
`
`(cid:2)
`
`Y(t) = k1
`U0K2
`
`ln
`
`1 + U0K2
`k1
`
`(1 − e
`
`−k1t)
`
`(cid:3)
`
`(2)
`
`ing force of the folding process they agree in the fact
`that a protein undergoes different more or less unstable
`conformations until it reaches its final native structure.
`At the absence of chaotropic agents these intermediates
`may exhibit intermolecular interactions, which leads
`to the major problems in a refolding procedure, aggre-
`gation and precipitation of the proteins. In refolding
`models estimating the final yield of a renaturation
`process, these competing side reactions are considered
`as of higher order while the folding reaction itself is
`approximated by a first order reaction (Zettlmeissl
`et al., 1979). A refolding reaction may therefore be
`described as
`= −(k1U + k2NU n)
`
`(1)
`
`dU
`dt
`
`with k1 the net rate constant of folding, k2 the net
`rate constant of aggregation, U the concentration of
`unfolded protein, t time, N aggregation number and n
`the reaction order of aggregation, assuming that back
`reaction from folded or aggregated protein to unfolded
`protein is negligible and formation of possible folding
`intermediates is infinitely fast. Analytical solutions of
`this differential equation exist for second (Kiefhaber
`et al., 1991) and third order aggregation reactions
`(Hevehan and De Bernardez Clark, 1997) and are
`depicted in Eqs. (2) and (3), respectively.
`
`where Y(t) is the yield of the refolding reaction, U0 the
`initial concentration of the denatured protein and K2 is
`the apparent rate constant of aggregation, combining
`aggregation number and rate constant of aggregation
`in k2N.
`Y(t) = Ψ
`
`(cid:4)
`
`−1 [(1 + Ψ 2) e2k2t − 1]1/2 − tan
`
`−1 Ψ
`
`tan
`
`(cid:5)
`
`(3)
`with Ψ = (k1/ k2U2
`0 )1/2, where k1 again is the net rate
`constant of folding, k2 net rate constant of aggregation
`and U0 is the initial concentration of unfolded protein.
`Common refolding techniques aim to inhibit these
`side reactions to enhance the final yield of correctly
`folded protein. Major attention has to be drawn to
`the chemical as well as physical environment during a
`refolding process since folding and aggregation kinet-
`ics are heavily influenced thereby. Kinetic constants of
`a refolding process are of importance for the design of
`operation parameters such as dilution rate, final protein
`concentration and refolding time. Once optimal refold-
`ing conditions are found, yield would be solely defined
`by protein concentration if an ideal dilution process
`could be applied, since aggregation is a concentration
`driven process. The determination of kinetic constants
`
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`Fig. 3. Time course of the formation of native protein and aggregates
`during a refolding reaction.
`
`knowledge of size distribution and density of the par-
`ticles to be separated (Taylor et al., 1986). Monitoring
`separation in disc stack centrifuges based on different
`spectrophotometric properties of inclusion bodies and
`cell debris (Jin et al., 1994) supports centrifugal process
`design. Separation may prove critical if cell fragments
`and inclusion bodies have similar sedimentation prop-
`erties. Pressure treatment for cell disintegration has to
`be optimized as has been studied in detail by Wong
`et al. (1997). They showed that repeated homogenizer
`passes resulted in better fractionation of inclusion
`bodies and cell debris leading to increased inclusion
`body purity. As a result of cell breakage, outer mem-
`brane components are released and may get adsorbed
`to the inclusion body surface (Hart et al., 1990). These
`compounds can be removed by several detergent
`washing steps, however, detergents sometimes cause
`problems
`in subsequent downstream processing
`and are therefore possibly avoided (Choe et al.,
`2006).
`Chemical extraction of inclusion body protein as
`an alternative to mechanical means poses both advan-
`tages and disadvantages. Extraction directly from
`fermentation broths shortcuts unit operation steps,
`however release of high molecular weight DNA leads
`to increased viscosity of the solution which may
`cause severe problems for the capture of the product.
`Additionally high amounts of host cell proteins are con-
`tained in the extract. These problems were partly solved
`in different approaches. Removal of DNA could be
`achieved by precipitation with spermine (Choe et al.,
`2002) and also cheaper DNA-precipitants with com-
`parable efficacy are available (Choe et al., 2006). A
`method for the selective extraction of recombinant pro-
`teins produced as inclusion bodies was described by
`Falconer et al. (1999) and successfully transferred to
`pilot scale. In a first step, membranes were permeabi-
`lized with a combination of urea and EDTA and host
`cell proteins were extracted, while inclusion bodies
`were kept insoluble by surface oxidation with the help
`of a disulfide bond promoting reagent. After removal
`of extracted compounds by diafiltration using a mem-
`brane with a high cut-off value, inclusion body protein
`was solubilized under chaotropic and reducing condi-
`tions. Compared with conventional extraction methods
`consisting of mechanical cell disruption, centrifugation
`steps and solubilization of inclusion bodies, similar
`protein extraction and purity could be reached. As a
`
`(4)
`
`may be accomplished by a direct fit of an appropri-
`ate refolding model such as shown in Eqs. (2) and (3)
`to data from dilution experiments at different protein
`concentration collected over refolding time until the
`endpoint is reached, or by an iterative approach when
`only data for one concentration are available over the
`whole time range, while endpoint data are available
`for different protein concentrations (Fig. 3). For a sec-
`ond order aggregation reaction yield at infinite time is
`described by
`Y = k1
`U0K2
`
`(cid:6)
`
`ln
`
`(cid:7)
`
`,
`
`1 + U0K2
`k1
`
`therefore kinetic constants can be easily extracted from
`the data set. However, kinetic constants heavily depend
`on refolding conditions and have to be determined for
`every buffer to be used.
`The required scale of a refolding process influences
`choice of a distinct methodology.
`
`3. Isolation of recombinant protein
`
`While in lab scale cell lysis is often performed
`enzymatically yielding in almost complete degradation
`of cell walls, the industrial means of inclusion body
`isolation is mechanical disruption of cells followed by
`centrifugation. After high pressure homogenization,
`which results in a suspension of cell debris and the
`product, sedimentation of inclusion bodies has to be
`performed. The design of a centrifugal process requires
`
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`
`591
`
`drawback this method is only feasible for proteins with
`a high cysteine content. In a different approach it was
`aimed to disintegrate cells while maintaining inclusion
`bodies insoluble by treatment with a combination of
`Triton X-100 and EDTA (Lee et al., 2004). This method
`should be applicable for a wider range of recombinant
`proteins and might constitute a considerable shortcut
`in processing.
`
`4. Determination of refolding conditions
`
`As a starting point of each refolding reaction, the
`kind of solubilization strategy has to be considered.
`In case of chemical extraction at high denaturant con-
`centration refolding procedures can directly be carried
`out. Starting from isolated inclusion bodies most
`techniques aim to reach complete unfolding, which is
`best accomplished by chaotropic agents such as guani-
`dinium chloride (GdnCl) or urea at high concentration
`in combination with reducing agents. Depending on
`the protein to be refolded higher final yield could be
`obtained by retaining certain native-like secondary
`structure already present within the inclusion bodies.
`This could be achieved by using detergents (Puri
`et al., 1992), buffers at high pH (Khan et al., 1998;
`Singh and Panda, 2005), GdnCl or arginine at low
`concentration (Tsumoto et al., 2003a) or even sodium
`hydroxide (Mahmoud et al., 1998; Suttnar et al., 1994)
`as solvent reagents. However this strategy is very
`protein dependent and has no general applicability. As
`a second step refolding has to be initiated by removal
`of denaturant and providing conditions which allow
`intramolecular interaction and formation of correct
`structure.
`To ensure optimal yield proper refolding conditions
`have to be found for every single protein. This is mostly
`done in an empirical approach based on former experi-
`ence. If little is known about the protein of interest, this
`may result in a vast number of different experiments
`since a large number of refolding additives have been
`described in the past (De Bernardez Clark, 1998). This
`includes denaturants in low concentration, polyols such
`as sugars or sugar alcohols, ionic or non-ionic deter-
`gents and organic solvents. They may either promote
`folding of the protein or inhibit aggregation. The influ-
`ence of GdnCl and l-arginine has been investigated in
`detail by Umetsu et al. (2003). They have studied the
`
`folding behavior of antibody fragments at the presence
`of GdnCl, l-arginine and redox systems in a stepwise
`dialysis system. Different chemical and spectroscopi-
`cal means were applied to determine aggregation, for-
`mation of structure, exposure of hydrophobic patches
`and formation of disulfide bonds. Effects of various
`detergents and organic solvents on refolding yield of
`lysozyme were investigated by Yasuda et al. (1998)
`to find cheap refolding additives allowing processing
`at high protein concentration. Formation of aggregates
`was monitored by dynamic light scattering. Again it has
`to be emphasized that a positive effect of an additive on
`refolding of a certain protein may cause the opposite
`for another one since protein properties are extremely
`diverse.
`In small scale refolding chaperones (Buchner et al.,
`1992), folding helper proteins, artificial chaperones
`(Machida et al., 2000; Rozema and Gellman, 1996) and
`redox pairs such as GSH-GSSG are used to improve
`yield. The use especially of chaperones is not practi-
`cal in large scale, since they have to be available in
`a stoichiometric proportion to ensure efficacy. How-
`ever, immobilization of folding aids allows a more
`efficient use of mostly expensive enzymes as shown
`with minichaperones (Altamirano et al., 1997), GroEL
`and GroES (Preston et al., 1999), oxidoreductases shuf-
`fling disulfide formation (Tsumoto et al., 2003b) or
`even a combination of chaperones (Altamirano et al.,
`1999). Reuse of artificial chaperones has been reported
`by Mannen et al. (2001). They used immobilized
`cyclodextrin to remove detergent from a denatured pro-
`tein to allow refolding. Operation of the process in
`circulating expanded bed mode allowed the stripping
`of detergent in a refolding requirement comparable to
`a batch suspension system but had the advantage of
`scalability.
`A fractional factorial design of the experimental set-
`up significantly reduces the effort (Tobbell et al., 2002).
`Additionally automation of screening allows a first
`evaluation of different refolding conditions. Vincentelli
`et al. (2004) describe an automated screening system
`based on the detection of precipitation of proteins by
`turbidity measurements in a 96 well plate format. They
`associate solubility of the protein to native structure.
`Data provide valuable clues in terms of additive selec-
`tion, however care has to be taken concerning soluble
`aggregates or stable misfolded species, therefore an
`activity assay is essential to draw further conclusions.
`
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`
`An adjacent optimization of the refolding buffer is
`indispensable. This includes evaluation of minimum
`additive concentrations providing a satisfactory yield
`as well as possible exchange of expensive buffer com-
`ponents to cheaper ones which is of specific concern for
`large scale applications. Additionally temperature, pH
`and of course redox potential may be crucial in a refold-
`ing reaction. Once proper conditions are found, optimal
`buffer exchange as well as protein concentration have
`to be evaluated.
`
`5. Refolding by dilution
`
`The simplest way to initiate refolding is to dilute
`the unfolded protein in refolding buffer. Both the con-
`centrations of chaotrops and proteins are decreased in
`a single step therefore permitting intramolecular but
`preventing intermolecular interaction. A final protein
`concentration of 10–100 ␮g/ml is generally applied in
`fast dilution procedures. It is the method of choice in
`industry because of simplicity of the process. Only a
`stirred tank and feeding pumps are required. In the sim-
`plest setup only temperature has to be controlled. The
`resolubilized protein from inclusion bodies is dispersed
`in the refolding buffer and the solution is kept for a
`fixed time, then the refolded protein is harvested. In
`large scale this technique poses several disadvantages
`despite of the simplicity of the processing scheme. For
`scale up of a stirred tank reactor mixing time should
`be kept constant. However this would be accompanied
`with an extreme increase in power input, most time,
`power per volume is kept constant, therefore mixing
`time increases with scale. Industrial scale devices have
`mixing times lasting several minutes (Doran, 1995).
`Uniform and fast mixing as required for the rapid
`dispersion of the feed stream is therefore hardly achiev-
`able with common mixing devices and formation of
`aggregates may evolve from local high protein con-
`centration due to the imperfect mixing. An advanced
`mixing reactor, the oscillatory flow reactor, was suc-
`cessfully applied for the refolding of lysozyme (Lee et
`al., 2002, 2001). In this device mixing intensity is cor-
`related to an oscillatory Reynolds number ReO, which
`is defined by reactor geometry, oscillatory frequency
`and amplitude,
`ReO = DωxO
`
`(5)
`
`ν
`
`where D is tube diameter, ω the angular frequency of the
`oscillator drive, xO the oscillatory amplitude and ν is the
`kinematic viscosity. The better scalability of the reactor
`in terms of mixing uniformity poses an advantage to a
`stirred tank reactor.
`Rendering dilution a continuous process in a com-
`parably small mixing device followed by a plug flow
`reactor (Terashima et al., 1996) might pose another
`answer to improved scalability and offers the possi-
`bility of a direct connection to capture systems such
`as expanded bed chromatography (Ferr´e et al., 2005).
`Additionally a continuous process allows the recycling
`of aggregated protein and therefore an increase of yield.
`However selection of operating parameters has to be
`optimized to retain productivity of a process as shown
`by Schlegl et al. (2005a,b). Using a continuous stirred
`tank reactor for refolding ultrafiltration devices have
`to be introduced for the removal of chaotrops and
`reducing agents evolving from the solubilization of
`inclusion bodies to retain a constant refolding environ-
`ment (Hohenblum et al., 2004; Schlegl et al., 2005a,b).
`A consequence of low protein concentration are
`large volumes to handle. Expensive refolding buffer
`supplements as mentioned above have to be provided in
`high quantity and wastewater treatment is another cost
`driving factor in the process. Concentration steps have
`to be included in the production scheme. This is either
`done by ultrafiltration or accomplished in subsequent
`purification steps such as ion exchange chromatogra-
`phy.
`Higher final protein concentration could be reached
`by pulse renaturation (Rudolph and Fischer, 1990) or
`fed batch dilution (Katoh et al., 1999). Applicability of
`these methods is based upon the stability of native pro-
`teins, which exhibit less or no tendency to aggregate
`compared to folding intermediates. In an optimized
`time scheme denatured protein is added to the reac-
`tion device. Intervals of protein addition or feed flow
`rate are dependent on protein folding kinetics. Addi-
`tion of protein is limited to the amount of denaturant
`which still allows refolding and has no negative effect
`on already folded protein.
`
`6. Pressure treatment
`
`An alternative strategy is to use pressurized tanks as
`refolding reactors which is especially valuable for pro-
`
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`
`593
`
`teins with a high tendency to aggregate during refolding
`or even during purification. Hydrostatic pressures of
`150–200 MPa permit the folding reaction while disfa-
`voring aggregation or completely reverse aggregation
`(Gorovits and Horowitz, 1998). Aggregates evolving
`from agitation, chaotrop-induced aggregates and bac-
`terial inclusion bodies were subjected to high pressure
`treatment at non-denaturing GdnCl-concentration (St.
`John et al., 1999). Even at protein concentrations up to
`8.7 mg/ml high recovery rates of native protein could be
`reached. Additionally pressure treatment of inclusion
`bodies for a longer period of time led to significant
`levels of active protein. P22 tailspike protein could
`be refolded from aggregates (Foguel et al., 1999).
`Extended studies of folding and aggregation at dif-
`ferent pressure conditions were carried out and it was
`shown that native protein was formed out of aggregates
`after pressure treatment even of short periods (Lefebvre
`et al., 2004). Therefore this could also be used as a
`recycling strategy in a continuous refolding process.
`Ligand binding domains of nuclear receptors, pro-
`duced as bacterial inclusion bodies, could successfully
`be activated by means of pressure refolding without the
`use of denaturants, proving that also proteins, which
`are prone to aggregation even in their native state can
`be refolded by this method (Schoner et al., 2005).
`The method permits refolding reactions at higher con-
`centration therefore reduces processing volumes and
`diminishes the need of chaotrops significantly.
`
`7. Large scale chromatographic refolding
`
`Many chromatographic refolding procedures have
`been described in the past as reviewed recently
`(Jungbauer et al., 2004). Size exclusion based as well
`as adsorption based chromatographic refolding pro-
`cedures have the advantage of operating at higher
`protein concentration compared to conventional dilu-
`tion. However applicability in a large scale process is
`often restricted. Especially size exclusion chromato-
`graphic refolding on a batch column has limitations due
`to small sample sizes and long run times. To achieve
`productivity of a process it is therefore inevitable
`to run it continuously. Schlegl et al. have character-
`ized a continuous matrix assisted refolding process of
`bovine ␣-lactalbumin (Schlegl et al., 2003) as well as
`of a recombinant therapeutic protein (Schlegl et al.,
`
`2005a,b) using annular chromatography at different
`operating conditions including recycling of aggregates.
`In case of the aggregation prone therapeutic protein sig-
`nificant increase in yield could be reached compared
`to a dilution process. Simulated moving bed (SMB)
`chromatography using four size exclusion columns was
`successfully applied for the renaturation of lysozyme
`as a model compound (Park et al., 2005). The process
`was designed using lysozyme and denaturant partition
`coefficients obtained from batch column experiments.
`Simulation of the process was in good agreement to
`obtained experimental results. However applicability
`of the process for the refolding of crude denatured pro-
`tein solutions remains to be shown. Sample application,
`which is critical for successful size exclusion chro-
`matography, might create problems when using crude
`extracts and especially the continuous regeneration
`of the chromatography matrix, which requires rather
`harsh conditions for the removal not only of unspe-
`cific bound protein aggregates but also other residual
`host cell compounds, poses a challenge to maintain the
`stability of the process.
`Expanded bed chromatography is suitable for deal-
`ing with crude samples. Capture of the target protein
`can be accomplished directly from cell homogenates
`as shown by Cho et al. (2001). They used an ion
`exchange matrix to adsorb a recombinant fusion
`protein, wash out cell debris and unbound components
`and exchange the buffer to initiate refolding, all in
`expanded bed mode. Stability of the bed could be
`maintained by restricting the applied sample volume
`therefore diluting the high molar urea buffer, which
`would have higher density than the used chromatog-
`raphy resin. Ion exchange in fixed bed mode has
`been described as a valuable tool for chromatographic
`refolding processes (Creighton, 1990; Li et al., 2002;
`Stempfer et al., 1996). A continuous process for the
`refolding of ␣-lactalbumin was described by Machold
`et al. (2005). Folded protein could be separated from
`aggregated protein, which was recycled therefore
`increasing the final yield of the reaction.
`
`8. Analysis of folded proteins
`
`Recombinant proteins used as therapeutics have
`to be proven to fulfill the required quality criteria.
`Most important is, besides high purity, the activity
`
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`
`of the compound which is closely correlated to its
`native structure. Activity assays combined with circu-
`lar dichroism spectroscopy are classified as adequate
`means to prove the identity of the product. Typically,
`CD spectra of unfolded, native and refolded protein
`are compared and the latter ones have to show equal
`features.
`The correct formation of disulfide bonds is fre-
`quently determined by reversed phase chromatography.
`Intermediate states, wrong as well as correct forma-
`tion of disulfide bonds can effectively be separated
`and monitoring the folding status during the process is
`therefore possible (Goldenberg and Creighton, 1984;
`Wu et al., 1998).
`Absence of aggregated protein has to be proven after
`purification of the refolded protein. This may be accom-
`plished by analytical size exclusion chromatography
`with on-line stray light detection. The intensity of scat-
`tered light increases with the size of a compound in
`solution, therefore aggregates create a large signal and
`even traces can be detected.
`
`9. Conclusions
`
`Albeit refolding by dilution is still the preferred
`technology for large scale refolding, a lot of alterna-
`tive strategies have been developed in the past. Most
`promising seem matrix assisted refolding using simple
`chromatography sorbents and pressure driven refold-
`ing. Still, the goal of future developments is to find
`methods to increase the final protein concentration
`under which refolding is performed.
`
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