[Advances in Biochemical Engineering/Biotechnology] Physiological Stress Responses in Bioprocesses Volume 89 || Analysis and Control of Proteolysis of Recombinant Proteins in Escherichia

Analysis and Control of Proteolysis of Recombinant Proteins in Escherichia coli

Aleksei Rozkov1 · Sven-Olof Enfors1

1 Department of Biotechnology, Royal Institute of Technology (KTH), Roslagstullsbacken 21,10691 Stockholm, Sweden E-mail: [email protected]

Abstract Proteolysis is one of the reasons for poor production of recombinant proteins inEscherichia coli. Important properties of E. coli proteases, which are relevant for the productionof recombinant proteins, are reviewed. Furthermore, various strategies to control the proteo-lysis of the recombinant proteins are presented. These strategies for control of proteolysis canbe applied on various stages of the process: design of more stable protein, a modification ofthe host cell in respect to proteolytic activity, optimisation of cultivation and downstream pro-cessing. However, before implementing these measures the proteolysis rate should be measuredin order to calculate a potential benefit of reduced proteolysis rate.

Keywords Proteolysis · Escherichia coli · Protease · Recombinant proteins

1 Role of Proteolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . 164

2 E. coli Proteases . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

2.1 Lon (La) Protease . . . . . . . . . . . . . . . . . . . . . . . . . . . 1662.2 ClpAP (Ti) Protease . . . . . . . . . . . . . . . . . . . . . . . . . . 1672.3 ClpYQ (HslUV) Protease . . . . . . . . . . . . . . . . . . . . . . . 1672.4 Proteases of the Cell Envelope . . . . . . . . . . . . . . . . . . . . . 168

3 Energy-Dependence . . . . . . . . . . . . . . . . . . . . . . . . . . 169

4 Susceptibility to Proteolysis . . . . . . . . . . . . . . . . . . . . . . 169

5 Impact of Proteolysis on the Yield of Recombinant Proteins . . . . 170

6 Measurements of Proteolysis . . . . . . . . . . . . . . . . . . . . . 171

7 Strategies to Control Proteolysis in E. coli . . . . . . . . . . . . . . 172

7.1 Control of Proteolysis on the Protein Level . . . . . . . . . . . . . . 1737.1.1 Sequence Modification . . . . . . . . . . . . . . . . . . . . . . . . 1737.1.2 Protective Fusion . . . . . . . . . . . . . . . . . . . . . . . . . . . 1737.1.3 Inclusion Body Formation Control . . . . . . . . . . . . . . . . . . 1737.2 Control of Proteolysis on Cell Level . . . . . . . . . . . . . . . . . . 1747.2.1 Use of Protease Mutations . . . . . . . . . . . . . . . . . . . . . . . 1747.2.2 Use of Host Strain Deficient in the Stringent Response . . . . . . . 175

© Springer-Verlag Berlin Heidelberg 2004

Adv Biochem Engin/Biotechnol (2004) 89: 163 –195DOI 10.1007/b95567

7.2.3 Co-Expression of Protease Inhibitors . . . . . . . . . . . . . . . . . 1757.2.4 Secretion to Periplasm . . . . . . . . . . . . . . . . . . . . . . . . . 1767.3 Control of Proteolysis on Cultivation Level . . . . . . . . . . . . . . 1767.3.1 Temperature Optimisation . . . . . . . . . . . . . . . . . . . . . . 1767.3.2 Optimisation of pH . . . . . . . . . . . . . . . . . . . . . . . . . . 1777.3.3 Addition of Protease Inhibitors to the Culture Medium . . . . . . . 1777.3.4 Use of Complete Medium or Supplementation of Amino Acids . . . 1777.3.5 Effects of Starvation and Extreme Growth Limitation

in High-Cell-Density Fed-Batch Cultures . . . . . . . . . . . . . . 1787.3.6 Influence of Toxic Metabolic Products . . . . . . . . . . . . . . . . 1807.3.7 Optimisation of Induction Strategy . . . . . . . . . . . . . . . . . . 1817.3.8 Control of Scale-Up-Specific Effects . . . . . . . . . . . . . . . . . 1827.4 Downstream Processing Level . . . . . . . . . . . . . . . . . . . . . 184

8 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . 185

9 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186

1Role of Proteolysis

Proteolysis in E. coli can be divided into two categories. The first is a proteolyticprocessing, which involves cleavage of a specific peptide bond to yield an activeor mature form of the protein. The second type of proteolysis is when the pro-tein is cleaved at multiple sites eventually resulting in its complete degradationto amino acids. Only this type of proteolysis, but not the proteolytic processing,will be covered in the present work.

There are two main physiological functions of the proteolytic degradation inE. coli. The first is inactivation of the short-lived regulatory proteins [1, 2], andthe second is a degradation of the unwanted, incorrectly synthesized, misfoldedproteins. These unwanted proteins are most likely devoid of biological functionand could even be toxic to the cell [3]. Recombinant proteins may also fall underthis category, often regarded by the cell as unwanted. Significance of proteolysiscan be judged from the fact that more than 3% of the enzymatic activities presentin E. coli at any given time are proteolytic [3]. Extracellular proteases are exportedto the medium and cleave proteins in order to provide cells with peptides and freeamino acids. In pathogenic bacteria these proteases can be virulence factors [4].Extracellular proteases are more common in Gram-positive bacteria and have notbeen discovered in E. coli.

Eukaryotes have a mechanism of marking proteins that are destined for degra-dation by attachment of the 8-kDa protein ubiquitin. The degradation itselfis taking place in specialized organelle called 26S proteasome, which shares com-mon traits with degradation by the ClpAP [5, 6] and ClpYQ [7] proteases inE. coli. Proteolysis in eukaryotes will not be covered here and the reader is refer-red to the following review articles [1, 6, 8–10].A broader review on proteases waspublished recently [11].

164 A. Rozkov · S.-O. Enfors

2E. coli Proteases

Proteolytic enzymes can be classified on the basis of many criteria: cellular location (cytoplasmic, periplasmic, membrane), energy dependency, nature ofthe active site (serine, aspartic, cystein and metalloproteases) etc. ([11]).Accord-ing to modern classification [12], depending on whether cleavage is taking place

Analysis and Control of Proteolysis of Recombinant Proteins in Escherichia coli 165

Table 1 Proteases in E. coli [3, 30, 50, 56, 67, 108, 146]

Protease Substrate ATP- Location Typedependency

Lon (La) SulA, RcsA, Yes Cytoplasm Serinelarge abnormal proteins

ClpAP (Ti, Clp) Large abnormal Yes Cytoplasm Serineproteins, MazE, sS

ClpXP lO, sS Yes Cytoplasm SerineHslUV (ClpYQ) SulA, RcsA, s32, Yes Cytoplasm Serine

large abnormal proteins

Fa Cytoplasm SerineSo Large proteins No Cytoplasm Serine

Oxidatively damaged proteins

Ci Small (<15 kDa) No Cytoplasm Metalloproteaseproteins

HflA Cytoplasm SerineFtsH (HflB) s32, lCII, SecY Yes Cytoplasmic Metalloprotease

membraneMi No Periplasm SerineTsp (Re, Prc) Oxidatively No Periplasma Serine

damaged proteinsRecombinant protein after cell disruption

HtrA (DegP, Do) Misfolded, abnormal No Periplasma SerineproteinsSecreted recombinant proteins

Pi (protease III) Small (<7 kDa) No Periplasm MetalloproteaseproteinsHighly sequence-specific betweentyr-leu, phe-tyr

OmpT Secreted recombinant No Outer Serine(protease VII) proteins membrane

a Previously protease HtrA was reported to be cytoplasmic and Tsp was reported as equally distributed between cytoplasmic and periplasmic fractions [49].

inside a protein or at its terminus, the proteases (also called ‘peptidases’) are clas-sified as endopeptidases (proteinases) or exopeptidases [13]. Adding to con-fusion, the term ‘peptidase’ is often used for those proteases that degrade oligo-peptides [12].

Most of the protein degradation in E. coli starts from initial cleavage by an energy-dependent protease as a rate-limiting step, followed by energy-indepen-dent proteases and peptidases [9], which degrade peptide fragments to freeamino acids [3, 14]. The first step must be controlled with a high precision in order to avoid indiscriminate proteolysis. An important distinction of energy-dependent proteases from those usually active in the extracellular medium, like,e.g. Bacillus proteases, is a quite large size of the former (up to 750 kDa) com-pared to 20–30 kDa of the latter. This difference probably reflects the presence ofstructural features of energy-dependent proteases that allow for more stringentsubstrate specificity and regulation [9]. The activity of peptidases must be veryhigh, since very little of intermediate products of proteolysis were found [9]. Foran overview of peptidases reader is referred to the review of Miller [15].

In the last two decades a number of E. coli proteases have been discovered.Most of them are summarized in Table 1.

The first-discovered and best-characterized E. coli proteases are Lon and Clp.Each of them constitutes about 0.05–0.2% of the cellular protein and togetherthey account for 70–80% of the energy-dependent degradation of proteins in vivo[3, 16].

2.1Lon (La) Protease

The Lon protease, also named La, degrades abnormal proteins, which may resultfrom nonsense or missense mutations, misincorporations or denaturation [9]and certain short-lived regulatory proteins [1]. The Lon protease exists in vivo as a tetramer (molecular weight 360 kDa) [17]. All monomers are essential forprotease activity despite the fact that each monomer molecule contains both ATPase and proteolytic sites. Although small peptides can be degraded withoutATP, it is required for degradation of large proteins [17]. The ATP role in degra-dation of large proteins is thought to be unfolding of the protein substrate andits translocation to the protected proteolytic chamber of Lon [18]. From two tofour ATP molecules are consumed per broken peptide bond under optimal conditions [3, 9], but this ratio increases to 13 if ADP is present and pH is not optimal [19]. Michaelis-Menten (Km) constant of Lon for ATP during proteolysishas been reported to be 0.20–0.25 mmol/l [20] and 0.02–0.05 mmol/l [21].ADP has a higher affinity towards Lon than ATP, thus completely preventing ATPbinding when both adenosine nucleotides are present in equimolar concentra-tions. This ADP inhibition is relieved considerably when a protease substrate ispresent [22, 24].

Lon generates polypeptides of 5 to 20 amino acids, which are degraded furtherby other endoproteases and peptidases without expenditures of ATP [19, 22–24].Therefore, the overall energy expenses on proteolysis are small, compared to theenergy for the cell growth. Lon is not essential to E. coli, but its deletion leads to


mucoidy (i.e. increased production of capsular polysaccharides) and increasedsensitivity to DNA-damaging agents [1]. An increased lon expression, which istaking place during the heat shock [25] or after the induction of recombinantproteins synthesis [9], leads to an elevated rate of proteolysis [26].

Degradation of some abnormal proteins by Lon requires the participation ofcertain heat shock proteins (DnaK, DnaJ, GrpE and GroEL), the deletion of whichsometimes have a higher effect on proteolysis than deletion of lon itself [27, 28].Possible mechanism of participation of these heat shock proteins is that they pre-sent abnormal proteins to proteases [29] or that they maintain the substrate in a unfolded conformation, which facilitates degradation [3, 30]. An alternative hypothesis exists that proteases and molecular chaperones just compete for thesame substrate, abnormal and unfolded proteins, without interaction with eachother [31].

2.2ClpAP (Ti) Protease

E. coli deficient in lon shows just 30% reduction in the ATP-dependent proteolysisof abnormal proteins [32], thus pointing on existence of other ATP-dependentproteases. ClpAP (Caseinolytic protease), also called Ti [33], is a large (750 kDa)complex consisting of one hexamer of ClpA subunits and one dodecamer of ClpPsubunits. ClpA (83 kDa) has an intrinsic ATPase activity, which increases in thepresence of ClpP and degradation substrates [1]. ClpP (21 kDa) is a heat shock-induced proteolytic subunit, which can, independently from ClpA, degrade smallhydrophobic peptides [34]. Degradation of large proteins requires their unfold-ing by ClpA and translocation into the ClpP chamber through a narrow (10 Å)channel [6, 35] with C-terminus entering first [36]. The chaperone function ofClpA appears to serve as a protection against indiscriminate degradation of cyto-plasmic proteins and probably determines the specificity of ClpAP protease. ClpAand ClpP have almost no homology to each other [1]. Similar to Lon, ClpAP generate polypeptides of about 1500 Da [9] and have almost the same energy requirements for proteolysis of large proteins (from 6 to 10 ATP molecules perpeptide bond) [1, 3] and Michaelis-Menten constant (Km) of ATP for proteolysis(0.21 mmol/l) [33]. Unlike Lon, ATP was shown to be necessary not only forproteolysis, but also for an assembly of the ClpAP complex.ADP inhibits both theformation of ClpAP complex [37] and proteolysis of the protein substrates. TheClpAP protease has a preference for a cleavage of peptide bonds after amino acidswith hydrophobic side chains [1].

2.3ClpYQ (HslUV) Protease

Double lon clp mutants still have 20–40% of the energy-dependent proteolysis of abnormal and recombinant proteins [38] and preserve almost all capacity ofstarvation-induced proteolysis [3, 16], which hinted on existence of at least oneunknown cytoplasmic ATP-dependent protease. The ClpYQ, also called HslUV,protease was discovered [39] and described to participate in the turnover of s32


and abnormal proteins [40]. ClpYQ (HslUV) similarly to ClpAP consists of wokinds of subunits: HslV – 19 kDa (250-kDa multimer 12–14 subunits) and HslU –50 kDa (450-kDa multimer 8–10 subunits) [41]. Michaelis-Menten constant (Km)of ATP for protein degradation is 0.3 mmol/l [41, 42]. Substrate ranges of Lon,ClpAP, ClpXP and ClpYQ are overlapping to some extent [35] and all these proteases have a homologous region of about 100 amino acids in the substrate-recognition domain [43].

Although ClpB is a homologue of ClpA, its deletion does not affect proteolysis.The action of ClpB in vivo is most likely to prevent or dissolve protein aggregates[30] and clpB-deficient strains have poor viability at high temperature [44].For more details about Clp protein family the reader is referred to several reviewarticles [5, 45–47].

2.4Proteases of the Cell Envelope

ATP has no stimulating effect on proteolysis when added to a periplasmic frac-tion of E. coli [48, 49], which is not surprising since there is no ATP in theperiplasm. The only ATP-dependent protease of the cellular envelope is FtsH(HflB), which is situated on the cytoplasmic membrane. FtsH has an ATPase domain at its C-terminus facing the cytoplasm and it appears to participate in diverse cellular activities [50, 51] including degradation of s32. FtsH is the onlyessential energy-dependent protease in E. coli [52].

HtrA (also called DegP and Do) is a periplasmic protease and a heat shockprotein regulated by the alternate heat shock sigma factor sE [53, 54]. HtrA actsas a chaperone at low temperatures refolding misfolded proteins, and as a pro-tease at elevated temperatures. Apparently, refolding is easier at lower tempera-tures and the proteolytic function of HtrA is used when refolding fails [55]. HtrAusually destroys unfolded, damaged or abnormal membrane or periplasmic pro-teins [3, 54] and has been reported to degrade secreted recombinant proteins[56–59]. HtrA is essential for growth at high temperature [53]. Overexpression ofHtrA was shown to reduce inclusion body formation and increase amount ofactive recombinant penicillin acylase in periplasm [60].

OmpT is an outer membrane protease, which could degrade secreted recom-binant proteins in vivo, in crude cell extract [3, 61] or even after protein refold-ing from inclusion bodies [62, 63]. The OmpT activity was found to increase inresponse to the induction of a recombinant cytoplasmic protein (CAT) [64, 78].A homologue of OmpT, OmpP, is involved in degradation of the endogenousmembrane protein SecY [65]. Prc (Tsp – Tail-specific protease) is a protease,which cleaves proteins at their C-termini after small hydrophobic residues suchas alanine and valine. Hydrophobicity of C-terminus increases the likelihood of cleavage [66]. Prc protease was implicated in degradation of the recombinantprotein FlgF after disruption of the cells [67]. DegQ and DegS are homologues ofthe DegP (HtrA, Do) protease [68]. DegQ is a periplasmic serine endopeptidase,which degrades casein and shares to some extent the substrate specificity withDegP. Deletion of degQ has no obvious growth defects, but degS is essential for thecell. DegS is apparently also a serine protease and is associated with cytoplasmicmembrane. Neither DegQ nor DegS are heat inducible.


3Energy-Dependence

Significance of the energy-dependent proteolysis for E. coli can be judged by thefact that it constitutes more than 90% of the overall protein degradation occur-ring in the cytoplasm [3]. Energy is not needed from point of view of thermo-dynamics, since a protein degradation by common proteases like trypsin is aspontaneous process and Lon and ClpAP do not require ATP for degradation ofsmall peptides [1]. Therefore, the energy (ATP)-dependency must serve to over-come some unfavourable step of reaction or to introduce an additional control.ATP hydrolysis appears to be needed either to change a conformation of the protease, or to unfold the protein substrate and to move the protein moleculethrough the active sites of the proteases [3, 6].

4Susceptibility to Proteolysis

The majority of proteins is stable, for exception of few small regulatory proteins(half time shorter than the cells doubling time) and abnormal proteins, whicharise from missense and nonsense mutations, amino-acid misincorporations,misfolded proteins, heat or chemically damaged proteins as well as some re-combinant proteins [1]. The picture becomes even more complicated when a pro-tein, normally stable, is misfolded and then targeted by proteases. This shows that target recognition depends not only on the primary structure, but also onsecondary and tertiary structure, i.e. degradation signals are either buried andnot accessible to proteases or should be formed from the damaged parts of theprotein [1]. In misfolded proteins these degradation signals somehow are pre-sented to the proteases.Accessibility of the cleavage site could be a reason for highsusceptibility of chimeric proteins for the cleavage at linkage sites, which oftenare exposed to the environment. The sequence that the protease recognizes mightnot be the one that is cleaved [69]. Also, it is possible that the protease must recognize several sequences at once [1].

Presence of certain amino acids (tryptophan, tyrosine, phenylalanine, arginineand lysine) in the N-terminus of a protein molecule is often shown to destabilizethe protein [70]. Known as the N-rule, this could explain why initial cleavage ofthe protein usually limits the overall degradation rate. Both Lon and Clp pro-teases tend to cut proteins in hydrophobic regions, thus yielding unstable frag-ments with hydrophobic amino acids at their termini [1], which are degradedeven faster.A similar rule was discovered for C-terminus. The presence of hydro-phobic amino acids in the last five positions was reported to destabilize the protein, while substitution of them with hydrophilic amino acids stabilized it[71]. This effect was greatest for the very last amino acid position. In anotherwork, an insertion of 1–3 copies of the tetrapeptide Ala-Trp-Trp-Pro close to theC-terminus of stable protein ZZT0 increased progressively its sensitivity to proteolysis [69]. “The C-rule” was shown to be involved in the degradation ofincompletely synthesized proteins by the ClpXP and ClpAP proteases. These proteins are tagged by SsrA peptide if the translation is stalled, thus marking


them for the subsequent degradation [72–74]. The proteases Prc (Tsp, whichstands for tail-specific protease) [66], and, possibly, HtrA, [54] were also shownto follow the C-rule.

5Impact of Proteolysis on the Yield of Recombinant Proteins

There could be many reasons for poor yield of recombinant proteins besides pro-teolysis. Therefore, before attempting to implement any measures to improveproduction by controlling proteolysis, the degradation rate of the product shouldbe measured and the potential benefits of reduced proteolysis assessed.As a rule,the rate of proteolysis during the process cannot be considered constant, there-fore measurements throughout the cultivation are necessary. The basis for cal-culations is the mass balance Eqs. (1) or (2), depending whether proteolysis hasfirst or zero order kinetics:

dP/dt = qp – kdeg · P – m · P, (1)or

dP/dt = qp – qdeg – m · P, (2)

where dP/dt is the accumulation rate of a protein with concentration P (mg/g),qp (mg/g/h) is a specific rate of synthesis, kdeg is a first order constant of prote-olysis (1/h), qdeg is a zero order proteolysis rate (mg/g/h) and m (1/h) is a cellgrowth. This equation was used for calculation of: 1) kdeg (or qdeg) in degradationtests with protein synthesis and bacterial growth inhibited by chloramphenicol,2) qP using kdeg obtained from degradation tests and 3) hypothetical accumula-tion rate, dP/dt, assuming a stable protein (kdeg is zero).

The proteolysis rate was determined in tests with inhibition by chloramphe-nicol [75]. The protein synthesis and cellular growth are inhibited, thus accumu-lation rate would reflect only proteolysis (Eqs. 3 and 4):

dP/dt = –qdeg (3)or

dP/dt = –kdegP. (4)

Once the proteolysis rate is determined, the synthesis rate of the protein product(qP) can be calculated. Then, in order to assess the potential gains in SpA yield,assuming all proteolysis is eliminated and neglecting an eventual negative effectof fast SpA accumulation and its high level in a cell, a hypothetical product ac-cumulation rate can be calculated. The simulation shows the potential for a con-siderable gain in the product accumulation (Fig. 1): the SpA concentration wouldreach over 400 mg/g at the end of cultivation compared to 14.7 mg/g obtained inthe experiment. In practice, however, these levels are unattainable because ofproduct inhibition. The fact that the protein SpA concentration was increasingonly during the first 3 h of induction does not mean stop of synthesis, since a highdegradation rate indicated that synthesis rate of SpA was maintained at the samehigh rate in order to sustain a constant level of SpA in the cells. It can be seen asa consequence of the first order kinetics since the rate of degradation increasestogether with substrate concentration.


6Measurements of Proteolysis

A number of methods is used for monitoring of proteolysis (reviewed also in[76]). The measurements of the mRNA transcripts of protease genes [64, 77, 78]could be a method of choice when studying the induction of proteases in re-sponse to fast changes in environment, for example, in heterogeneous environ-ment of large-scale bioreactor [64, 77, 78]. Analysis of protease proteins them-selves (using 2D gel electrophoresis [79] or Western blotting) can be used for thesame purpose if a short time resolution is not an issue. The amount of mRNA oreven the proteases themselves does not necessarily correlate with protease ac-tivity. The technique called SDS-GPAGE [80], where the first G stands for gelatin,enables to monitor activity of proteases towards model substrate gelatin. Thistechnique is based on conventional SDS-PAGE with addition to a gel of gelatin,which later serves as substrate during incubation of a gel in a buffer, which contains necessary co-factors for protease activity. The proteolytic activity resultsin degradation of gelatin, which is later visualized during staining [80]. The overall proteolytic activity can also be measured by the rate of release of radio-actively labelled matter from cells resuspended in unlabelled medium (pulse-and-chase) [81]. In many cases the rate of proteolysis is sometimes judged by thepresence and amount of degradation bands on SDS-PAGE gels. Because of theprocessive nature of E. coli ATP-dependent proteases [3] and the fast degradationof peptide fragments by peptidases this “method” has little value for quantifica-tion of proteolysis [76]. In order to measure the proteolysis rate of specific pro-teins the samples from pulse-and-chase experiment must be separated on 2D gel


Fig. 1 Intracellular concentrations of a truncated staphylococcal protein A (SpA) (squares) andthe stable fusion protein SpA-gal (circles) during fed-batch cultivations performed underidentical conditions. Dashed line represents theoretical SpA concentration calculated assum-ing no degradation and the synthesis rate calculated from Eq. (1). Arrow indicates the point ofinduction. (Data from Enzyme Microbiol Technol (2000) 27:743–748)

electrophoresis, proteins spots of interest quantified and compared. Alternativetechnique involves incubation of cells in the glucose-supplemented mineralmedium with chloramphenicol (100 mg/l) to inhibit protein synthesis with subsequent analysis of samples by Western blotting [75, 82, 83]. The membranesare scanned, bands of the product integrated and the result is plotted againsttime. The slope of the protein concentration represents the degradation rate of this product (Fig. 2). Depending on the level of the protein product in the cell, the proteolysis may have different order kinetics. Before or immediately after induction when protein concentration in the cell is low compared to that ofproteases, proteolysis is substrate-limited and has a first-order kinetics. Whenprotein is accumulated to higher concentration the degradation may become enzyme- (protease) limited and, therefore, zero-order in relation to the pro-tein. The example of changed proteolysis kinetics during a cultivation is given in Fig. 2.

7Strategies to Control Proteolysis in E. coli

Strategies for control of proteolysis were reviewed earlier in the following articles:[116, 217]. Other aspects of the recombinant protein production are covered indepth elsewhere [63, 84–89].


Fig. 2 Changed kinetics of proteolysis from first order at low product concentration to zero order at high product concentrations. Cells were taken from beginning (0.5 h after induction,circles) and the end of ZZT2 production (8.5 h after induction, squares) and incubated withchloramphenicol to inhibit protein synthesis. Aliquots of the culture were then subjected to Western Blot, scanned and remaining protein concentration is plotted (Fig. 5 from EnzymeMicrobiol Technol (2000) 27:743–748)

7.1Control of Proteolysis on the Protein Level

7.1.1Sequence Modification

If a linker between the domains of the fusion protein contains amino acid sequence recognized by proteases, it could particularly be vulnerable to degra-dation due to its exposure to the environment. If this amino acid sequence is notessential for the function of the protein it can be deleted or modified.A success-ful example of application of this strategy was described [90], when the fusionprotein SpA-bgal, which was originally degraded by the OmpT protease at a basicdipeptide sequence in the linker, was stabilized after removal of the sensitive site.

Other concerns, which should be addressed in designing a plasmid for proteinproduction, although not directly related to proteolysis, are the metabolic burdencaused by the expression of genes coding for antibiotic resistance [85] and thetoxicity of polylinker-encoded peptides [91].

7.1.2Protective Fusion

Numerous examples [92] show that a protein may be stabilized by fusion to otherprotein or even to a copy of itself. The comparison of accumulation levels ofproteolytically unstable protein SpA and the derived from it fusion protein SpA-b-galactosidase is shown in Fig. 1. The results showed that the accumulationrate of SpA-bgal was much higher compared to SpA and the final intracellularconcentration of SpA-bgal reached 138 mg/g, compared to only 14.7 mg/g for SpA(Fig. 1). The fact that the SpA-bgalactosidase level did not reach the theoreticalpredicted level shows that accumulation of this protein was limited by other factors. Interestingly, the position where the fusion partner is attached also in-fluences the susceptibility to degradation. The VP1 protein of the foot-and-mouth disease virus joined to the C-terminus of b-galactosidase was sensitive to proteolysis, while the same protein at the N-terminus was stable [93]. Evenmore efficient protection from degradation can be achieved if the recombinantprotein is protected by fusion partners from both termini (“dual affinity fusionstrategy”) [94].

7.1.3Inclusion Body Formation Control

Formation of inclusion bodies usually stabilizes proteolytically sensitive proteinsdue to the steric hindrance for proteases, although limited proteolysis of aggre-gated protein in inclusion bodies may still occur [95, 96]. The decision whetherto direct protein design and fermentation conditions towards formation ofinclusion bodies depends on many factors, such as proteolytic stability of theproduct, feasibility of refolding and the cost considerations. However, if the protein is stable in the cytoplasm, the formation of inclusion bodies should be


avoided, since refolding may be complicated and expensive process with a littlefinal yield. Different methods to reduce inclusion body formation exist, both ongenetic level, like fusion to solubilizing partner like Z-domains [97, 98], thio-redoxin [99, 100], E. coli NusA protein [101] or maltose-binding protein [102],or on cultivation level based on the regulation of the rate of synthesis of the protein with temperature [84, 103, 104], partial induction (reviewed by Donovanwith co-workers [86]) or addition of non-metabolizable carbon source such asdesoxyglucose [105]. Too high protein synthesis rate is considered to be a pri-mary reason for inclusion body formation for cytoplasmically localized proteins[105], and, therefore, a slower synthesis rate usually favours formation of solubleprotein [104, 106, 107]. Other strategies include a co-expression of the molecularchaperones to overcome the folding pathway limitations, in the cytoplasm[108–110], as well as in the periplasm [104]. Molecular chaperones can also be induced by addition heat-shock-inducing agents like ethanol (3%) [225].The reader is referred to following review articles for more detail [84, 92, 105,107, 111].

7.2Control of Proteolysis on Cell Level

7.2.1Use of Protease Mutations

A straightforward solution to reduce the proteolysis could be to delete the genescoding for responsible proteases. Although the ATP-dependent proteases, whichcause most of the damage to production of recombinant proteins: Lon, ClpAPand ClpYQ are not essential for the cell, their deletions reduce the strain fitness.Kanemori et al. found that the triple protease mutant (DhslVU-clpPX-lon), whileenabling to improve the stability of recombinant protein fivefold, had a specificgrowth rate half as low as the parent strain and was not able to grow at 42 °C [40].Moreover, a ClpP-deficient strain in an other work [83] also resulted in aboutthreefold increase of specific product concentration, but also had a lower biomassyield and higher cell lysis when grown in high-cell-density fed-batch cultures.E. coli deficient in Lon protease has a distinctive mucoid and UV sensitive pheno-type, which is a consequence of the stabilization of two Lon substrates: RcsA, apositive regulator of transcription of genes involved in capsule production(cps–capsule synthesis genes) [32, 112, 113], and SulA, an inhibitor of cell division[114, 115]. Mucoid phenotype can be overcome by either growth conditions (e.g. use of rich media or temperatures higher than 37 °C) or certain secondarymutations, like inactivation of gal and cps genes [116]. Filamentation is higher in rich medium, especially with yeast extract. Deletion of sulA does not have adeleterious effect on cell growth, and, therefore, can be used to control fila-mentation [116].

Instead of introducing the protease mutations some researchers choose towork with strains naturally devoid for some proteases, like BL21 strain, whichlacks Lon and OmpT proteases. Corchero et al., for example, used it to obtain ahigher yield of recombinant fusion protein [93]. In the other case, the BL21 strain


was used for a production of rat neuronal nitric oxide synthase.When expressedin the other, OmpT+ Lon+ E. coli strain this protein was heavily degraded uponcell lysis [117], suggesting that OmpT was responsible for proteolysis. The use ofthe BL21 strain of E. coli has another advantage – a lower accumulation of acetatewhen grown on glucose [118].

The double mutant for cell envelope proteases DegP and Tsp could be grownto high cell densities (50 g/l) in fed-batch cultures, yielding 4.5 times more ofsecreted fusion protein A-b-lactamase compared to the reference culture [119].Moreover, the triple mutant for cell envelope proteases OmpT, DegP, and proteaseIII increased production of this protein more than sixfold [120], which was seenas promising. However, the mutations for envelope proteases also had some negative effects. The tsp mutant was reported to have an increased permeabilityof outer membrane [121]. Although the quadruple mutant for all known cell envelope proteases except for FtsH (DdegP-ptr-ompT-tsp) exhibited a reductionof maximum specific growth rate in the LB medium only by 17% compared toreference strain, it grew poorly in a defined medium [119]. In other cases, DegP-deficient strains were successfully applied for the production of the cytochromesubunits FixO and FixP from B. japonicum [122], diphtheria toxin [58] and enterotoxin mutant proteins [59].

Deletion of the htpR (rpoH) gene coding for s32 transcription factor may reduce rate of proteolysis even more than lon mutation [116, 123], since all cyto-plasmic ATP-dependent proteases are upregulated by the heat shock. Moreover,combined htpR and degP deletion had a much more dramatic effect on accumu-lation of secreted recombinant fusion protein A-blactamase than in either ofthe single (DdegP or DrpoH165) mutants [124]. This result may implicate the pro-tease FtsH, which is under regulation of s32 [125], in the remaining proteolyticactivity towards protein A-blactamase.

7.2.2Use of Host Strain Deficient in the Stringent Response

Dedhia and co-workers [126] reported that the use of relaxed strain of E. coli(DspoT-relA), enabled one to improve production of chloramphenicol acetyl-transferase (CAT) in fed-batch culture up to 20-fold due to avoidance of stringentresponse together with concomitant increase of proteolytic activity and transla-tional inhibition, which were usually observed during production of CAT protein[80, 127–131].

7.2.3Co-Expression of Protease Inhibitors

It has been observed that an infection with T4, T5 and T7 phages inhibits degra-dation of abnormal proteins, but does not affect degradation of normal proteins[132, 133]. The expression of the gene pinA from T4 in E. coli was shown to improve the yield of recombinant protein human fibroblast interferon [134],selectively inhibiting the Lon protease by interfering with its ATPase function[135, 136]. PinA had no effect on other ATP-dependent proteases ClpAP or ClpYQ


or ATP-independent proteases (trypsin, chymotrypsin, subtilisin and pepsin).E. coli lon+ cells that produced protein pinA have phenotype of lon mutants andthe lon mutation had no additional effect in the cells producing PinA protein.Protease FtsH can be inhibited by lcIII protein [137]; however, since FtsH is essential, this phenotype is lethal. Numerous serine protease inhibitors wereidentified from mammals, birds, reptiles, plants and micro-organisms. The best-characterized example of such inhibitors is ecotin, an E. coli periplasmic protein,which is a powerful inhibitor of a wide range of trypsin-like proteases. The mech-anism of action is thought to be substrate-like binding of ecotin to the active siteof protease [138].

7.2.4Secretion to Periplasm

One efficient strategy to reduce proteolysis is to fuse a signal peptide to N-endof the protein in order to enable secretion of the protein to periplasm. Althoughsome proteolysis is still taking place in the periplasm, different kinds of proteasesare active there (Table 1). Secretion to periplasm enabled to increase the stabil-ity of preproinsulin tenfold [139]. Besides reduction of proteolysis secretion toperiplasm has also other advantages, like disulfide bond formation and simpli-fied recovery of the protein.

7.3Control of Proteolysis on Cultivation Level

7.3.1Temperature Optimisation

The temperature can influence degradation of recombinant proteins in severalways. First, protein degradation will decrease with decreased temperature in accordance with the Arrhenius Law, as was demonstrated in a case of un-stable recombinant b-galactosidase [140]. Second, protein misfolding oftenoccurs at elevated temperatures due to either direct heat damage to the proteinor due to an increase of recombinant protein synthesis rate, which leads to alimitation in a folding pathway. This may render a protein, which was recognizedas native under normal conditions, more susceptible for proteolysis. In this case lowering the temperature of cultivation may prevent this. However,decreased temperature will not likely prevent proteolysis if the protein isdegraded in its folded native form. In this case misfolding and subsequentaggregation into inclusion bodies might be even beneficial to prevent degrada-tion. Finally, transcription of genes coding for periplasmic HtrA protease and all cytoplasmic ATP-dependent proteases is under regulation of s32 or sE

transcription factors which are induced at elevated temperatures. This effectmight be a reason behind additional increase of proteolysis rate of unstable b-galactosidase at temperatures higher than 40 °C [140]. Successful examples oftemperature optimisation include production of interferon a-2 [141] and 6-phos-phofructo-2-kinase [142].


7.3.2Optimisation of pH

The E. coli periplasm is permeable to relatively small molecules and, therefore,must have a pH equal to or at least close to the extracellular pH. With exceptionof OmpT, all cell-envelope and periplasmic proteases have pH optimum underalkaline conditions and are inhibited at pH lower than 6.0. This enabled to control proteolysis of secreted recombinant protein simply by changing pH to5.5–6.0, where concentration of recombinant protein A-b-lactamase was maximaland about 4-fold higher than at pH 7.0 [120]. An inhibition of protein synthesiswith chloramphenicol confirmed that a reduction of proteolysis rate was a reasonfor this increase of protein A-b-lactamase concentration. Since the cytoplasmicpH is tightly regulated [143], any attempt to shift it will probably result in an increase of the maintenance energy requirements or (and) reduced ATP pool,which could negatively affect protein synthesis and upregulate proteolysis.

7.3.3Addition of Protease Inhibitors to the Culture Medium

A number of serine protease inhibitors, including diisopropyl fluorophosphate,the sulfonyl fluorides, tosyl lysine chloromethyl ketone, and the aromatic di-amines prevent increase of proteolysis of normal proteins during starvation fora carbon or nitrogen source. However, these inhibitors failed to decrease the basallevel of proteolysis in the growing cells and degradation of abnormal proteins[144]. These findings suggest the existence of two proteolytic systems in E. coli.One of them is responsible for degradation of normal proteins during growthand degradation of abnormal, and possibly recombinant proteins during allphases of culture. This type of proteolysis is not inhibited in vivo by the inhibitorsmentioned above. The second type of proteolysis is the one stimulated by star-vation and could be inhibited by the protease inhibitors [144]. OmpT [145] andpossibly another unidentified cell envelope protease [56] has shown to be inhibitedby zinc ions. Metalloproteases are inhibited by EDTA [146] due to chelating metalion necessary for protease function.

7.3.4Use of Complete Medium or Supplementation of Amino Acids

Overexpression of proteins is a burden for cell metabolism due to the competi-tion with a synthesis of normal proteins for ribosomes, charged tRNAs and precursors. This problem may become worse due to an unusual amino acid com-position of the recombinant protein, which can result in a temporary depletionof this amino acid pool and induction of the stringent response. Feeding ofphenylalanine, which is overrepresented in the recombinant protein chloram-phenicolacetyl transferase (CAT), was shown to alleviate the stress caused by thisproteins overexpression. This enabled one to increase the yield of CAT presum-ably due to reduction of proteolysis [129]. Addition of amino acids, which are present in biomass and recombinant protein in equal amounts did not improve


the yield, while increasing concentration of the overrepresented amino acid evenresulted in a lower CAT concentration, due to feedback inhibition of biosynthesisof not only this amino acid, but also of those sharing common biosynthetic pathways [128]. Besides the loss of yield, a high rate of synthesis of recombi-nant proteins with unusual amino acid composition can lead to amino acid misincorporations [147], which can be partially overcome by the use of richmedium [148].

Numerous pieces of evidence show that the use of rich medium enables to improve the production of recombinant proteins. Tsai et al. detected no re-combinant protein, human IGF-1, if the cells were grown on minimal medium.However, if yeast extract and bactotryptone were used the specific yield ofIGF-1 increased to 30 mg/g [149]. Specific yield of recombinant malaria antigenincreased from 15 to 22 mg/g when complete medium was used instead ofminimal medium [150]. Addition of casamino acids to the culture producing b-galactosidase under trp promoter enabled to increase the specific yield ofthe product twofold [151]. A specific yield of recombinant lipoprotein antigenagainst Lyme disease was improved in a rich medium (Super Broth) from 4- to30-fold dependent on the temperature [152].

7.3.5Effects of Starvation and Extreme Growth Limitation in High-Cell-Density Fed-Batch Cultures

The overwhelming majority of E. coli proteins have a half-life significantly higherthan a doubling time for exception of few regulatory proteins [153]. Proteolysisrate of the bulk proteins in E. coli depends on genetic background and environ-mental conditions. In the growing cells protein turnover is about 1% per hour,which increases to 5% [3] when cells are starved for ammonia, carbon, aminoacids [154] or inorganic nutrients [155].Apparently the purpose of this increasedrate of proteolysis is to provide amino acids for the synthesis of the so-called star-vation proteins, necessary for the cells to survive in the new conditions [156, 157].Cells potential to survive starvation is greatly compromised by the inhibition ofprotein synthesis during first hours of starvation [158]. Starvation for glucose,which stimulates twofold increase of proteolysis is accompanied by a moderate(30–50%) decrease in an ATP pool [159, 160]. This stimulation of proteolysis wasobserved only for normal cellular proteins, but not for abnormal ones, which arealready degraded at a very high rate. Apparently, degradation of the abnormalproteins has the highest priority for the cell, which commits its last energy resources for their degradation. When the ATP pool was decreased to 5–10% ofthe normal value, the proteolysis of all proteins: normal, abnormal [160] or recombinant [82], was inhibited. The starvation itself was not necessary for stimulation of proteolysis of normal proteins, as was shown in the experimentswith stringent and relaxed strains of E. coli and specific inhibitor of ppGpp syn-thesis. Proteolysis of normal proteins was found to be proportional to ppGppconcentration. Moreover, changes in ppGpp concentration was followed byinstantaneous (less than 2 min) adjustment of the proteolysis rate, thus exclud-ing the possibility of synthesizing new proteases de novo. Two explanations were


proposed to explain the ppGpp effect for stimulation of proteolysis: i) reversibleactivation of existing proteases by ppGpp; ii) ppGpp renders proteins more sus-ceptible to proteolysis [161].At elevated temperatures (42 °C) proteolysis increasewas no longer dependent on ppGpp [159], since at this high temperature the rea-son for increased proteolysis is probably an appearance of misfolded proteins.Early observations that a major part of starvation-stimulated proteolysis wasinhibited by chloramphenicol [162] were first interpreted as if the increase ofproteolysis during starvation required the protein synthesis. However, St. Johnand co-workers [159] explained the inhibiting effect of chloramphenicol onstarvation-induced proteolysis by the accumulation of charged tRNA, which istaking place due to the inhibition of protein synthesis and subsequent decreaseof ppGpp levels. Starvation-induced proteolysis is ATP-dependent [32], but theprotease which is responsible for protein degradation during starvation has notbeen identified, and no evidence exists so far that new proteases are synthesizedduring starvation. Although lon– mutation dramatically stabilizes abnormalproteins it does not affect starvation-induced proteolysis [32, 144, 163].

Most of the work on stress responses was carried out with starving cells atquite low cell densities. However, relatively little is known whether this knowledgecould be extrapolated to the fed-batch cultures, which are usually used for pro-duction of the recombinant proteins. Limited oxygen and heat transfer, especiallyin the large-scale reactors [164], necessitate growing cells at very low specific ratein order to achieve a high cell density. These conditions are usually associatedwith changes in cell physiology like elevated levels of ppGpp, loss by cells ofability to divide [165, 166], induction of ss-dependent genes [167], filamentation[168], decrease in mRNA stability [169] and segregation of cells into sub-popu-lations with cytoplasmic membrane depolarization, and cell death [170].Although reverse correlation was observed between ppGpp levels and specificgrowth rate during glucose limitation in both fed-batch and A-stat [171] cultures,the most dramatic increase in ppGpp and ss was observed during transition frombatch to fed-batch culture, i.e. during transition from unlimited to limited growth[166, 172]. Besides inhibiting protein and RNA synthesis [173] the stringent re-sponse is known to increase proteolysis, as was mentioned earlier, either directly[161] or by positive regulation of the Lon protease [174]. Moreover, elevated tran-script levels of nine genes were detected in high-cell-density fed-batch cultureproducing recombinant protein CAT [226], among them genes involved in DNArepair (recA and uvrB), proteolysis (degP and groEL) and cell lysis (mltB). How-ever, caution should be exercised when interpreting a decline in protein concen-tration as increase of proteolysis [168, 175], since this may rather be attributedto a reduction of the rate of synthesis, while proteolysis rate stays constant or evendeclines [83, 176]. To our knowledge, no evidence exists so far to claim that thegrowth limitation in fed-batch cultures induces an increased rate of proteolysis,which could be expected in case of starvation [154, 177]. The induction of thestringent response could also be an indicator of the stress caused by a high syn-thesis rate of recombinant protein. The ppGpp concentration in E. coli cells wasshown to correlate with the rate of synthesis of recombinant proteins [178]. Sincethe amino acids obtained as a result of proteolysis may be used again for proteinsynthesis, this would relieve the metabolic burden on the cells. For this reason a


better measure of metabolic burden might be the accumulation rate of theproteolytically unstable protein product or even the ratio between the accumu-lation rate and cellular growth rate (which shows a relative burden of recombi-nant protein production on the cellular metabolism), primary structure andcodon usage of the protein product.

7.3.6Influence of Toxic Metabolic Products

Formation of by-products of glucose by overflow metabolism or mixed-acidfermentation is considered to be a problem in production of recombinant pro-teins. It is mostly acetate, which received most attention, since it accumulates inlargest concentration. Inhibition of the growth rate of E. coli W3110 was observedalready at acetate concentration as low as 1.3 g/l [179], but the biomass yield decreased only when acetate concentration was 2.5 g/l (by 16%) and 5 g/l (by24%). Inhibition of synthesis of recombinant protein was observed at acetateconcentrations higher than 4 g/l [103] or 10 g/l [180]. The mechanism of acetateinhibition is probably related to its action as uncoupler of membrane potential[181, 182], which will increase maintenance energy requirements or reduce intra-cellular pH [183]. It should be noted that the inhibition by acetate is stronger at low pH due to the higher ratio of undissociated form of acetic acid, which cantravel freely across the cytoplasmic membrane. Other possible reasons for poorerproduction of recombinant proteins with concomitant production of acetatecould be related to a repression of TCA cycle enzymes when E. coli is grown onrich medium supplemented with glucose [184] or low energy status of the cellsin anaerobic conditions [185].

Acetate formation can be reduced by many methods [186], one of the mostcommon of them being a use of fed-batch culture in order to limit biomassgrowth rate. Although this technique enables to avoid acetate formed by over-flow metabolism, the formation of acetate and some other metabolic products by mixed-acid fermentation is more difficult to control. Glucose hetero-geneities common in the large-scale bioreactors often lead to a local high con-sumption of oxygen, thus creating zones of high glucose and low oxygen con-centrations [187], which favour formation of acetate, formate, lactate, succinateand ethanol.Although these metabolic products are easily re-consumed in well-mixed and aerated part of the bioreactor, formate has the slowest rate of re-up-take and, therefore, can accumulate in a culture broth [187]. Formate can also beformed in well-mixed bioreactors due to a poor diffusion of oxygen to the cellscoated with DNA, which is released as a result of biomass lysis [188]. Little isknown about the influence of formate on bacterial growth and metabolism,but it has been reported to inhibit DNA synthesis and cell division [189–192] andit is likely to share the property of other weak organic acids to decouple protongradient across cytoplasmic membrane. Decoupling of membrane potential mayresult in decline of energy status, which could increase the rate of proteolysis[159, 160]. Another concern during extended fed-batch cultivations is an accu-mulation of inorganic ions due to pH titration. In a study where sodium sulfatewas added to the culture the inhibition effect was already noticeable if concen-


tration exceeded 0.2 mol/l [103]. The negative effects of high salt and acetateconcentrations were additive and a possible underlying principle for this couldbe increase in the maintenance requirement and subsequent energy limita-tion [103].

7.3.7Optimisation of Induction Strategy

Very high expression level of recombinant proteins was reported to have manynegative effects on the cell: inhibition of growth, ribosome destruction [193],periplasmic leakage, cell lysis [194, 195], induction of SOS [196] and heat shock[197, 198] responses and other [199, 200]. These negative effects are observedeven if the protein itself is not toxic or have no obvious biological activity [201].These effects of the recombinant protein’s production constitute “metabolicburden/load”, which was defined by Glick [85] as “the portion of a host cell’s resources – either in the form of energy such as ATP or GTP, or raw materials asamino acids – that is required to maintain and express foreign DNA, as eitherRNA or protein, in the cell”. First, the use of strong promoters (e.g. T7) results inhigh quantity of recombinant mRNA, which competes with native mRNA for ribosome-binding sites, thus undermining synthesis of vital native proteins essential for cell viability and growth. The second reason is that the synthesis ofrecombinant proteins requires energy and metabolic precursors, which wouldotherwise be used for the synthesis of cellular proteins. An unusual amino acidcomposition and codon usage of overexpressed recombinant protein may also invoke stress responses. It was shown that the induction of recombinant proteinwith unusual amino acid composition could lead to an increase of proteolysis andpossibly an induction of new proteases [130]. This effect is thought to be relatedto the stringent response [178], when production of recombinant protein imposesa burden on the protein synthesizing machinery draining the pools of certainamino acids. Since overdesign is not likely in the biology world, the abundanceof tRNA species is proportional to the frequency of codon usage [202]. Expres-sion of a recombinant protein with rare codons may lead to frameshifts, aminoacid misincorporations [203] or premature termination of the translation [204].Whether this may lead to the increased proteolysis remains to be seen, but thechanges in protein quality will probably be more important than the loss of yielddue to the degradation. Three strategies used to overcome this problem are [87]:i) overexpression of lacking tRNA [205], ii) replacement of rare codons with morecommon ones, iii) use of a eukaryotic host. The effects of rare codons on production of recombinant proteins are reviewed elsewhere [203, 204]. A main-tenance of the recombinant plasmids also contributes to metabolic burden (re-viewed by Glick [85]), which, however does not constitute as large part as tran-scription and translation [165, 166]. The synthesis of additional protein fromantibiotic marker gene is often overlooked as an additional factor contributingto metabolic burden [85].

A decreased synthesis rate of recombinant proteins can be beneficial to thecells since this relieves a metabolic burden to the cells [178], thus reducing pro-teolysis and improving recombinant protein yield [127, 129]. Reduced synthesis


rate would also benefit the secretion and folding of recombinant proteins (seesection Inclusion Body Formation Control).

7.3.8Control of Scale-Up-Specific Effects

The usual scale-up criteria of fermentation are similar energy dissipation ratesor impeller tip speeds, or both [206]. This makes inevitable increase of mixingtime during scale-up [206]. While in laboratory and small pilot scale (less than50 l) bioreactors the mixing time is in the range of a few seconds [207], it in-creases dramatically in a larger scale. For instance, mixing time in a bioreactorwith a working volume of 8 m3 increases to 10–25 s (depending on power input)and in a 22-m3 culture to 150–200 s [208]. For some substances, which have a con-centration in a culture broth close to limiting (glucose, oxygen), their mixing timein the large-scale bioreactor could be longer than the rate constant of their con-sumption, which will inevitably lead to a formation of gradients. Such limitingsubstrates can be glucose, since it is usually used to control growth rate in fed-batch cultures and oxygen, because of its low solubility in water. Glucose gra-dients exist due to insufficient mixing of the concentrated (500 g/l) glucose feeding solution added to the culture broth where the typical bulk glucose con-centration is in a range of tens of milligrams per liter. Two types of glucoseheterogeneities exist in large-scale reactors: firstly, short term fluctuations ofglucose concentration in time, as revealed by rapid and frequent sampling froma specific point in the bioreactor [209, 210] and, secondly, spatial variation ofaverage (quasi steady-state) glucose concentration in different parts of the bio-reactor [187, 209, 210]. It has been established by the example of Saccharomycescerevisiae that microorganisms can respond to an shift-up of glucose in 2 s bychanges in intracellular metabolite pools [211]. An excess of glucose in the PFRsection of a scale-down reactor (Fig. 3) was shown to create oxygen limitation dueto the increased uptake of oxygen favouring the formation of formic acid andother mixed-acid fermentation products [187].As a result, these fluctuations withrespect to oxygen and glucose concentrations could influence the production ofrecombinant proteins because of the induction of the various related stress responses. Experiments with pulsed feeding in the well mixed lab-scale bio-reactor and in the scale-down reactor (Fig. 3) showed that the concentration of ppGpp increases dramatically within one minute after the consumption ofthe residual glucose [212]. Moreover, Schweder and co-workers showed by mRNAmeasurements that stress-related genes, proU, uspA and dnaK, are expressed inthe high glucose zone of the scale-down reactor [77]. proU is a gene encoding for a binding-protein-dependent transporter of glycine and l-proline, which is induced after an increase of osmolarity [213]. Osmotic shock may increaseproteolysis, possibly by induction of an osmodependent protease [214]. Thisincrease in osmolarity can also lead to elevated levels of the ss subunit of RNApolymerase [215], which governs the expression of stationary phase proteins.The heat shock protein DnaK has been suggested to participate in protein degra-dation [29] and its induction by low oxygen and high glucose may thereforeincrease the proteolysis of recombinant protein. Fermentation scale-up was


shown to have a significant impact on the production and proteolysis of recom-binant protein ZZT2 [176]. The final accumulation level of the protein productdeclined to 35 mg/g of biomass in the large-scale compared to 50 mg/g level inthe lab-scale, which was explained by a combination of a higher proteolysis rateand declining synthesis rate of the protein product in large-scale cultivation[176]. Reviews of mixing-related scale-up effects on physiology of E. coli andS. cerevisiae can be found elsewhere [216, 217].

Another type of heterogeneity in large-scale bioreactors is pH gradients dueto titration, which may have an impact on some processes [218]. Retrofitting of large-scale bioreactors with upward-pumping stirrers was found to be an efficient way to reduce compartmentalization [208, 219].

At last, an oxidative damage can occur in large-scale due to three reasons:i) possible use of oxygen-enriched air to overcome oxygen mass-transfer prob-lems, ii) high liquid head pressure in tall bioreactors, iii) high head space pres-sure to maintain sterility [220]. E. coli cells may respond to oxidative damage in many ways [220], including the necessity to dispose off the damaged proteinsby proteolysis.

Another challenge in fermentation scale-up is a limited oxygen and heat transfer capacity in large-scale bioreactors, which imposes an additional con-


Fig. 3 Principle of the scale-down reactor for analysis of physiological responses in a zone ofhigh glucose concentration. The plot shows schematically the concentration fluctuations with respect to glucose and DOT experienced by an individual cell circulating between the STRand PFR

straint on the maximum feeding rate [164]. Besides possible induction of stressresponses elicited at very low specific growth rate, a low feeding rate leads to a decreased cell yield due to the higher ratio of substrate used for the maintenanceenergy compared to the substrate used for biosynthesis.

7.4Downstream Processing Level

Use of protease inhibitors is a standard method to reduce proteolysis duringdownstream processing. Since there is no concern for inhibition of cellulargrowth and protein synthesis the inhibitors can be used more liberally comparedto the cultivation stage. Numerous inhibitors of serine proteases exist, e.g. diiso-propylfluorophosphate (DFP) [33, 221], phenylmethanesulfonyl fluoride (PMSF)[144]. Less toxic alternatives to serine protease inhibitors mentioned above are APMSF (4-amidino-phenyl-methane-sulfonyl fluoride) [222], AEBSF (4-(2-aminoethyl)-benzenesulfonyl-fluoride, hydrochloride) and sodium bisulfite(5–10 mmol/l), which also inhibits some aspartic proteases [223]. Many metallo-proteases are inhibited by EDTA (1–2 mmol/l), which chelates the metal ions inthe active centre of the protease. Many other products are available commercially,both individual proteases inhibitors as well as “protease inhibitor cocktails”. Pro-tease inhibitors are covered in depth in the following book [224].

It is a common knowledge that the rate of chemical reactions decreases withdecreasing temperature in accordance to Arrhenius kinetics. Therefore, if the culture broth is kept at 20 °C instead of 42 °C, where proteolysis is fastest, the remaining amount of the recombinant protein A will be about 2.5-fold higher after an hour after stop of de novo synthesis. At 4 °C almost all protein becomesstable (95%), while at 42 °C only 20% remained intact [75]. Cooling the culturequickly enough in a large scale can be problematic because of high ratio ofvolume to the area of heat transfer. If temperature is raised to higher than 55 °Cdegradation of protein A is decreased dramatically due to protease inactivation[75]. This technique is suitable for use if the product itself is heat-stable. Sinceheat inactivates the proteases irreversibly, the cells need to be exposed to the hightemperature only for a brief period time in a separate unit, which capacity shouldbe enough to treat all culture broth in a reasonable time.

The harvest of the product in large-scale may continue for hours, dependingon the capacity of the downstream equipment (cell separation, disintegration, gelfiltration etc.). Therefore if a protein is proteolytically unstable, it is important to optimise the conditions in the bioreactor during the transition period fromcultivation to downstream processing. Rozkov et al. studied the use of energy depletion in the cells to control proteolysis of a cytoplasmic recombinant protein.Removal of only oxygen or only glucose had little effect on proteolysis due to ATPgeneration from fermentation or respiration, respectively. Absence of glucosecombined with addition of sulfite was shown to reduce the ATP pool by 90% andto stabilize the recombinant protein A.Cell separation produced a similar stabiliza-tion effect due to the extremely high cell densities in the cell concentrate. Cell dis-ruption caused an additional decline in the ATP concentration due to the dilutionof ATP and action of ATPases, which also stabilized the protein A [82] (Fig. 4).


The disruption of cells may result in an exposure of proteins to proteases fromother cellular compartments. Use of fast protease (or product) separation couldbe advantageous in the cases where the cytoplasmic recombinant protein is degraded by ATP-independent proteases such as OmpT or Prc. Disruption of cellsin the presence of 8 M urea enabled one to prevent proteolysis of the recombi-nant FlgF protein [67].

8Concluding Remarks

Proteolysis may have a significant impact on production of recombinant pro-teins. In many cases the degradation of the protein product is not recognized asa reason for its low accumulated level because no or weak degradation bands arevisible on SDS-PAGE electrophoresis. This emphasizes a need for a proper mea-suring of proteolysis rate. In spite of its utmost importance, still little is knownabout the mechanism of the recognition of a recombinant proteins as a proteasetarget. Engineering of the recombinant protein, which is devoid of proteolyticallysensitive motifs or structures would be an efficient way to circumvent the problem.

As was shown, a number of techniques for control of proteolysis is available atevery stage of the process, usefulness of whose are still assessed by trial and error.Only a few studies exist where several techniques were compared to each other orused simultaneously. Little is known about influence of cultivation conditions, es-pecially in the fed-batch cultures, on proteolysis of recombinant proteins.

Acknowledgements This work was supported by the Swedish Research Council for EngineeringSciences (TFR) and by the Biotechnology program of the European Community (BIO4-CT95-0028).


Fig. 4 ATP levels in E. coli W3110 pRIT2 pRITcI857 during transition phase from cultivationto downstream processing. Filled bars represent the stages where ATP is too low to support energy-dependent proteolysis

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Received: December 2003


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[Advances in Biochemical Engineering/Biotechnology] Physiological Stress Responses in Bioprocesses Volume 89 || Analysis and Control of Proteolysis of Recombinant Proteins in Escherichia