CEA, Département d'Ingénierie et d'Etudes des Protéines, Centre d'Etudes de Saclay, Bât. 152, 91191 Gif-sur-Yvette, France
1 To whom correspondence should be addressed. E-mail: belin{at}dsvidf.cea.fr
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Abstract |
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Keywords: Escherichia coli/protein expression/recombinant antibody/selection system/toxicity
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Introduction |
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Among these different ways of optimizing recombinant protein production, bacterial strain engineering appears promising. Bothmann and Plückthun recently presented another approach of cellular engineering, not based on the rational tailoring of strains, but on the selection of strains improved for functional protein production (Bothmann and Plückthun, 1998, 2000
). They developed a phage display-based selection procedure to detect E.coli genes whose overexpression would increase the presentation of antibody fragments on the phage surface. They thus identified two E.coli genes, skp and fkpA, and demonstrated that their overexpression significantly increases the production of functional antibody fragments on the phage surface but also in the periplasmic space (
1.2- to 20-fold).
We were interested in developing this type of approach further and we wondered if it would be possible to select strains with chromosomal mutations that would be advantageous in functional protein production. The general idea was to screen rapidly a large number of mutated bacteria for the production of a functional protein. We based our strategy on the observation that aggregation of proteins in bacteria could be toxic to the cell. This is particularly true for proteins that are targeted to the bacterial envelope where aggregates are often found to be associated with membranes whose integrity they disturb (Bowden and Georgiou, 1990; Skerra and Plückthun, 1991
; Knappik et al., 1993
; Baneyx, 1999
). We reasoned that bacterial strains with mutations that would diminish aggregation in favour of correct folding of recombinant protein should display a less pronounced toxicity and hence could be selected in a toxicity assay. Such an approach, i.e. the selection of chromosomal mutations that reduce toxicity, has been widely utilized by geneticists to study essential pathways of E.coli, like exportation or envelope stress response (Shuman and Silhavy, 2003
). In the case of recombinant proteins, very few reports using this approach have been published despite the premier work of Miroux and Walker describing the selection of BL21(DE3) derivatives overexpressing toxic proteins (Miroux and Walker, 1996
). However, the strains selected by Miroux and Walker apparently displayed a reduced toxicity because of a reduction of the toxicity of the T7 expression system used in their study and not because of an increased production of functional recombinant protein (Miroux and Walker, 1996
).
As a model protein to overproduce, we chose a recombinant single-chain antibody fragment targeted to the periplasmic space. Production of recombinant antibody in the periplasmic space is often impaired by a toxicity characterized by limited growth and cell lysis (Plückthun and Skerra, 1989; Somerville et al., 1994
; Knappik and Plückthun, 1995
). Furthermore, it was observed that recombinant antibodies that give higher levels of production of functional molecules are less toxic to the bacteria (Forsberg et al., 1997
; Strachan et al., 1999
). This suggests the possibility of selecting bacteria producing more functional recombinant antibody, assuming a simple toxicity assay is available. However, the main problem with selection procedures based on reduction of protein toxicity is the frequent appearance of clones presenting a low level of expression of the toxic protein. In order to eliminate these false-positive clones, we combined our selection procedure with a phenotypic screen. To this end, we expressed recombinant antibodies as an N-terminal fusion to bacterial alkaline phosphatase PhoA, an enzyme whose activity can be easily monitored on plates by addition of a chromogenic substrate. By means of this scFv-PhoA recombinant antibody, we isolated and characterized chromosomal mutations advantageous for functional scFv-PhoA production.
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Materials and methods |
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Bacterial strains, phages and microbiological techniques
Strain BW25113 (F lacIq rrnBT14 lacZWJ16 hsdR514
araBADAH33
rhaBADLD78; from Mary Berlyn) was used for the constructions of the chromosomal deletions (Datsenko and Wanner, 2000
). Strains MC1061 [F araD139
(ara-leu)7696 relA1 spoT1 galE15 galK16
(lac)X74 rpsL hsdR2 mcrA mcrB1], W3110 [F inv(rrnB-rrnE)] and MC4100 [F araD139
(argF-lac)U169 rpsL150 relA1 flbB5301 deoC1 ptsF25 rbsR] were used for the production of recombinant antibodies. Strain LE392 (F hsdR514 glnV44 supF58 lacY1 galK2 galT22 metB1 trpR55) was used for the growth of phage
. Strain XL1-Blue [F'::Tn10 proA+B+ lacIq
(lacZ)M15/recA1 endA1 gyrA96 thi hsdR17 glnV44 relA1 lac; from Stratagene] was used for cloning experiments.
Bacteria were grown on LuriaBertani medium or SOC medium (Miller, 1992). Antibiotics were added at the following concentrations: ampicillin, 200 µg/ml; chloramphenicol, 25 µg/ml; kanamycin, 30 µg/ml; spectinomycin, 100 µg/ml.
Addition of 40 µg/ml 5-bromo-4-chloro-3-indolyl phosphate (XP) to the agar plate was used for the detection of alkaline phosphatase activity.
Transduction with P1 was done according to Miller (1992).
Phage 1105 (b221 cI857::Tnmini-kan Oam29 Pam80 nin5) was from Nancy Kleckner and was manipulated as described for the integration of the mini-kan transposon into the E.coli chromosome (Way et al., 1984
). The insertion site of the transposon was determined as previously described (Manoil, 2000
). DNA fragments, including the chromosometransposon junction, were amplified with two PCR steps. The degenerate primers used were the same as those described by Manoil. The primer hybridizing on DNA corresponding to the inserted mini-kan transposon sequence was 5'-ACTGATGAATGTTCCGTTGC-3'. The PCR fragments obtained were gel purified and the insertion site was determined by sequencing.
Gene disruption
Directed gene inactivation was carried out essentially as described by Datsenko and Wanner and consists in the replacement of part of the gene to be inactivated by a DNA fragment encoding resistance to an antibiotic. This is achieved by transforming bacteria expressing the Red recombinase system with PCR products suited for gene replacement and selected bacteria expressing the antibiotic resistance (Datsenko and Wanner, 2000
). PCR products were obtained by amplification of the chloramphenicol acetyl transferase (cat) gene carried on plasmid pKD3 using the Expand High Fidelity PCR System (Roche Diagnostics). The oligonucleotides used for this amplification are composed of a priming site region hybridizing on pKD3 (GTGTAGGCTGGAGCTGC-3' for the forward primer and CATATGAATATCCTCCTTAG-3' for the reverse primer) preceded by a region homologous to the 50 bp preceding the replaced region in the wild-type gene for the forward primer and the 50 bp following the replaced region in the wild-type gene for the reverse primer (see Table I). PCR products were first precipitated, digested with DpnI and then purified on agarose gel using the QIAquick Gel Extraction Kit (Qiagen). Two hundred to 1000 ng of PCR product was used to transform freshly made electrocompetent cells BW25113 carrying plasmid pKD46 that encodes the
Red recombinase system under the control of araB promoter. Transformants were selected as described. The disruption of the gene was verified by PCR on colony using oligonucleotides flanking the disrupted gene and sequencing the PCR product. The corresponding allele was then transduced to MC1061 with P1. The presence of the disrupted gene in the transductants was checked by PCR on colonies. The nomenclature used to describe the different alleles is the same as that described by Datsenko and Wanner (2000)
. According to the homologous extensions of the primers used, yheS was deleted from nucleotide 91 to 1550, wecB from nucleotide 51 to 830 and wecE from nucleotide 98 to 1101 and the mutations obtained were respectively named DE(yheS)1460::cat, DE(wecB)779::cat and DE(wecE) 1004::cat.
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In the figures, null mutations in yheS, wecB and wecE are named, respectively, DE(yheS), DE(wecB) and DE(wecE), independently of the presence or absence of the cat gene.
Plasmid constructions
All the enzymes used in this study were from New England Biolabs unless otherwise stated. DNA manipulations were done according to Sambrook et al. (1989).
Soluble recombinant antibody was expressed with plasmid pLIP5v. This plasmid derives from pLIP4 (Gillet et al., 1993) and is designed to construct hybrid protein with a potent variant of alkaline phosphatase, PhoAGN, whose expression is under the control of the Ptac promoter (Boulain et al., 1995
; Muller et al., 2001
). Plasmid pLIP5v-scFv17F12 carries the origin of replication from plasmid p15A, confers resistance to ampicillin and was used for the production of the hybrid protein scFv17F12-PhoAGN (Boulain et al., 1995
; Carrier et al., 1995
). Plasmid pLIP5v-scFvM
2-3 was used for the production of scFvM
2-3-PhoAGN hybrid protein. It was obtained by cloning between the SalI and SacI sites of pLIP5v a SalISacI DNA fragment obtained from pLIP5-scFvM
2-3 (Merienne et al., 1997
).
Plasmid pLIP5v-scFv17F12QUAD encodes a modified scFv17F12-PhoAGN hybrid with the residues F10, F73, L83 and F87 of the svFv changed to S10, L73, A83 and Y87, respectively. It was obtained by site-directed mutagenesis of pLIP5v-scFv17F12 using the QuikChange Site-Directed Mutagenesis Kit (Stratagene). These mutations were designed according to previously published work on antibody fragment framework engineering for efficient production in E.coli (Knappik and Plückthun, 1995; Ulrich et al. 1995
; Nieba et al., 1997
; Wall and Plückthun, 1999
).
Plasmid pLIP5-scFv17F12(His)6 was constructed to assure the production in the periplasmic space of scFv17F12 tagged with six histidine residues to its C-terminus. It was obtained by inserting a DNA fragment encoding six histidine residues and a stop codon in the SacI site of pLIP5v-scFv17F12. This cassette was made by annealing oligonucleotides 5'-CATCACCATCACCATCACTAATAGAGCT-3' and 5'-CTATTAGTGATGGTGATGGTGATGAGCT-3'.
DNA fragments encoding skp and fkpA were amplified from the DNA from E.coli strain W3110 by using Taq DNA polymerase and oligonucleotides 5'-CTTAGCCTTAGCT-GGATGCTGGATCTCGAGTAAGTGTTCTCCACAAAGGAATGT-3' and 5'-AATGTTCGGTAGATTACCTCATCTCGAGTCCAACTGCTGCGCTAAATCAG-3' for skp, 5'-CTTAGCCTTAGCTGGATGCTGGATCTCGAGGATTCACCTCTTTTGTCGAATGGT-3' and 5'-AATGTTCGGTAGATTACCTCATCTCGAGGGCTCATTAATGATGCGGGTAACTA-3' for fkpA, thus creating XhoI sites (in bold) flanking the amplified gene. DNA fragments were then cloned into pETBlue-1 vector by using the pETBlue-1 AccepTorTM Vector Kit (Novagen). These plasmids were digested with XhoI and the fragments encoding skp and fkpA were then cloned into the XhoI site of pLIP5v-scFv17F12.
All the constructions were verified by DNA sequencing using the ABI Prism 310 Genetic Analyser (Applied Biosystems).
Expression of recombinant antibodies and preparation of bacterial extracts
Bacterial strains were transformed with the corresponding plasmid and plated on freshly made LB agar medium containing 0.5% glucose and the correct antibiotic. After 24 h at 30°C, several colonies were picked up to inoculate a 2 ml culture of the same medium. After overnight incubation at 30°C, 25 ml of pre-warmed LB plus the appropriate antibiotic were inoculated with 500 µl of the overnight culture. Bacteria were then grown at 30°C until A600 reached 1.11.2, isopropyl ß-D-1-thiogalactopyranoside (IPTG) was added to 10 µM and the incubation was continued for 17 h before pelleting cells.
Periplasmic extracts were prepared as follows. Bacterial cultures (25 ml) were centrifuged in the cold and then gently resuspended in 6 ml of ice-cold 30 mM Tris pH 8, 20% sucrose and 0.5 mM EDTA. Lysozyme was added (final concentration 100 µg/ml) and bacteria were kept in the cold on a rotary shaker for 45 min. After centrifugation, the supernatant was saved and filtered through 0.45 µm. Protein concentration in the extracts was determined using the Coomassie Protein Assay Reagent Kit (Perbio Science) and was 0.250.30 µg/ml unless otherwise indicated.
Alternatively, bacteria were sonicated or passed through an Eaton pressure cell (Rassant). Bacterial cultures (25 ml) were centrifuged in the cold and the pellet was resuspended in 5 ml of phosphate buffered saline (PBS: 10 mM phosphate, 150 mM NaCl, pH 7.2)0.5 mM EDTA. One millilitre of this suspension was sonicated (2400 J over a 4 min period) while 4 ml were passed two times through an Eaton pressure cell. Bacterial extracts were then centrifuged for 20 min at 4000 g. Pellets were respectively resuspended in 1 (sonicated suspensions) or 4 ml (pressed suspensions) of PBS0.5 mM EDTA (referred to as low centrifugation pellets). Supernatants were then centrifuged in a Beckman 70.1 Ti rotor at 40 000 r.p.m. for 90 min at 4°C. Supernatants were saved as soluble fractions. Pellets were resuspended as before in PBS0.5 mM EDTA (referred to as high centrifugation pellets). Protein concentration in the soluble fractions was determined using the Coomassie Protein Assay Reagent Kit and was 2.12.4 µg/ml for soluble fractions obtained after sonication and 2.72.9 µg/ml for soluble fractions obtained after passage through the Eaton pressure cell. Protein concentration in the insoluble fractions was determined using the Dc Protein Assay (Bio-Rad) and was 0.670.96 µg/ml for insoluble fractions obtained after sonication and 0.370.51 µg/ml for insoluble fractions obtained after passage through the Eaton pressure cell.
SDSPAGE and western blot
SDSPAGE was performed according to standard procedures (Sambrook et al., 1989). Proteins were then transferred to nitrocellulose membrane (Optitran BA-S 83; Schleicher & Schuell) using the Mini Trans-Blot cell (Bio-Rad) and following the instructions of the manufacturer. Membranes were saturated overnight in PBS2% BSA. They were then incubated for 2 h with mouse monoclonal antibodies to bacterial alkaline phosphatase (1/2500 in PBS0.1% Tween-20; Caltag Laboratories), washed three times over 30 min with PBS0.1% Tween-20, and incubated for 2 h with peroxidase-conjugated F(ab')2 fragment goat anti-mouse IgG (1/5000 in PBS0.1% Tween; Jackson ImmunoResearch). After three washes with PBS0.1% Tween-20 over 1 h, staining was done with 0.4 mg/ml 3,3'-diaminobenzidine in 100 mM Tris pH 7.6 and 0.15% hydrogen peroxide.
ELISA
For the conjugate antibody fragments, the production was quantified by direct ELISA. Wells of MaxisorpTM microtitre plates (Nunc) were coated for 2 h at room temperature with 100 µl of antigen solution (10 µg/ml in 100 mM Tris pH 7.4). For scFv17F12-PhoAGN, human IgG were obtained from Sigma; for scFvM2-3-PhoAGN, toxin
from Naja nigricollis was obtained from J.-C.Boulain. Plates were saturated overnight with 200 µl of 100 mM Tris pH 7.4, 0.3% BSA. Serial dilutions (100 µl) of crude periplasmic extract in 100 mM Tris pH 7.4, 0.1% BSA, 1 mM MgCl2 and 20 µM ZnCl2 were added. After 2 h at room temperature, plates were washed three times with 10 mM Tris pH 7.4, 0.1% Tween-20. Values were read at 410 nm after addition of 200 µl of 1 M diethanolamine, 1 mM MgCl2, 20 µM ZnCl2 and 10 mM p-nitrophenyl-phosphate and standing for 1020 min at room temperature.
The production of scFv17F12(His)6 was quantified in periplasmic extracts by direct ELISA. Wells of MaxisorpTM microtitre plates were coated for 2 h at room temperature with 100 µl of 10 µg/ml human IgG solution in PBS. Plates were then saturated overnight with 300 µl of PBS1% BSA and washed five times with PBS0.1% Tween-20. Serial dilutions (100 µl) of crude periplasmic extracts in 0.1% BSAPBS were added and plates were incubated for 2 h at room temperature. After five washes as above, 100 µl of anti-(His)6-peroxidase antibody solution (Roche; 1/500 dilution in blocking buffer) was added to each well for 1 h at room temperature. Wells were washed as previously and detection was initiated by adding 100 µl of 0.1 M sodium acetate pH 4.3, 0.56 mg/ml 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) and 0.009% H2O2. Values were obtained by reading the absorbance at 410 nm after 10 min at room temperature.
Each point was done in duplicate. Each value presented in the figures is the mean of the two measurements. Each experiment was done two or three times and the results obtained were identical. A typical experiment is presented for each figure. Addition of 1 µg of soluble antigen per well of the 1/128 dilution reduced the signals to 0.10.15 in all cases.
The level of expression was assessed by comparing for two different extracts the fold dilutions necessary to obtain the same absorbance of 0.5 at 410 nm.
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Results |
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As a recombinant antibody model, we used a poorly expressed scFv that derives from the mouse monoclonal antibody 17F12 directed against human IgGs (Carrier et al., 1995). Plasmid pLIP5v-scFv17F12 encoding the scFv17F12-PhoAGN hybrid was used to transform different bacterial strains. In a first assay, bacteria carrying pLIP5v-scFv17F12 were grown overnight on LB agar plates without inducer and then isolated on the same medium containing XP and different concentrations of IPTG. Plates were incubated overnight at 30 or 37°C. For each temperature, we observed different growth rates according to IPTG concentration. However, they were not sufficiently pronounced to be transposable to a selection procedure. We then reasoned that bacteria might have more difficulty in coping with toxicity if it occurs during the transformation procedure. We then modified the procedure used for the transformation by spreading the transformation mix on LB agar plates containing directly different concentrations of the inducer IPTG in addition to XP and ampicillin. Plates were incubated overnight at 30 or 37°C, respectively. In this experiment, we observed an inhibition of growth relative to IPTG concentration. The best results were obtained with transformants of strain MC1061 incubated at 30°C. When grown in the presence of 100 µM IPTG, bacteria gave colonies homogeneous in size (1 mm diameter), deep blue and no growth inhibition was observed. On 200 µM IPTG, the majority of transformants grew poorly and gave very small colonies. Pale blue or white full-size colonies appeared (
5% of the number of colonies obtained on 100 µM IPTG) that represented mutants with reduced expression level. We then repeated the experiment with the plasmid pLIP5v-scFv17F12QUAD encoding an engineered scFv17F12-PhoAGN (see Materials and methods). The modification of the scFv resulted in a
400-fold increase in functional recombinant antibody recovered from the periplasm (see later). In this case, no difference was observed when transformants were spread on 100 or 200 µM IPTG: growth was identical to that observed without IPTG. These results indicate that the toxicity observed on plates can be related to the level of production of functional recombinant antibody, suggesting that our screen might be appropriate for the selection of clones with increased capacity to produce functional recombinant antibodies.
Isolation of mutants
The goal of this work was to demonstrate that it is possible to rapidly isolate chromosomal mutations allowing higher production of functional recombinant protein. With this objective in mind, the choice of the mutagenesis method was important because we needed to identify rapidly the chromosomal mutation present in the selected clones and to demonstrate that this mutation was responsible for the observed effects. We found that mutagenesis by transposon insertion into the chromosome was the only method that met our requirements. First, this method is rapid and efficient without the need for mutagenesis optimization. Secondly, the insertion site of a transposon can be easily determined with two PCR steps (Manoil, 2000). Thirdly, the effects of a transposon insertion can be verified by site-directed gene disruption (Datsenko and Wanner, 2000
). We therefore constructed a library of E.coli null mutants by using
1105 to deliver a mini-kan transposon into the chromosome of strain MC1061 (Way et al., 1984
). Approximately 10 000 kanamycin-resistant colonies were obtained. Bacteria from these colonies were pooled, made competent and transformed with plasmid pLIP5v-scFv17F12. Using our selection procedure, we isolated eight deep blue colonies and 21 pale blue colonies growing on 200 µM IPTG. Bacteria were then streaked on LB agar plates without antibiotic containing XP and 500 µM IPTG in order to cure the plasmid. The 29 clones gave white colonies that were further tested as ampicillin-sensitive, indicating the loss of the plasmid.
The bacteria of the 29 selected clones were then individually transformed with pLIP5v-scFv17F12 and tested for the periplasmic expression of functional scFv17F12-PhoAGN. The periplasmic extracts were prepared and tested by ELISA as described in Materials and methods. We found that eight clones out of 29 gave higher ELISA signals (data not shown). These eight clones corresponded to the deep blue clones selected, indicating a good correlation between phenotype observed on plates and production of functional recombinant antibody.
The transposon insertions of these eight clones were transduced with P1 into MC1061, and the transductants were tested for the production of scFv17F12-PhoAGN in the periplasmic extracts. The same 2-fold increase compared to the parental strain was obtained except for the insertion 2.3 that gave the same result as MC1061 (data not shown). The clone carrying the insertion 2.3 possibly accumulated one or several mutations unlinked to the transposon and allowing a reduced toxicity. The seven other clones for which the transposon insertions were responsible for the increase in antibody production were further analysed.
Inactivation of yheS, wecB or wecE induces an increase in recombinant antibody fragment production in the periplasm
The chromosomal insertion sites of the mini-kan transposons were determined as described in Materials and methods. The clones 8.1, 9.1, 11.1, 13.2 and 21.3 were found to carry different insertions in yheS, a gene of unknown function encoding a putative cytoplasmic protein (located at min. 75 of the E.coli physical map; Rudd, 1998). The clones 4.1 and 5.1 carry transposon insertion in the genes wecB and wecE, respectively. These genes belong to a gene cluster (min. 85.5 of the E.coli physical map) involved in the synthesis of the enterobacterial common antigen (ECA), a cell surface glycolipid (Reeves et al., 1996
; Rick and Silver, 1996
).
In order to check that inactivation of each of these genes was responsible for the increased production observed, we constructed by homologous recombination a directed chromosomal disruption of wecB, wecE and yheS according to Datsenko and Wanner (2000). The different mutants carrying these disruptions were tested for the production of functional scFv17F12-PhoAGN in the periplasm (Figure 1A). For each mutant strain, we observed an
1.5-fold increase compared to the wild-type strain, thus confirming that cassette inactivation of wecB, wecE or yheS induces an increased production of the functional recombinant scFv17F12-PhoAGN in the periplasmic extract.
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To discount the possibility that the effects observed with the different null mutations are linked to the bacterial strain MC1061 used for the screening and the production, we tested the production of functional antibody fragment in another E.coli genetic background, by using strain W3110 (Figure 2). We observed a 2- to 3-fold increase in production of functional recombinant scFv for the mutant strains. These results show that the three null mutations selected with our screening procedure can be used in another genetic background. Furthermore, a more significant increase may also be obtained, as observed for the yheS mutation which induces a 3-fold increase in recombinant scFv-PhoAGN production in W3110 versus a 1.5- to 2-fold increase in MC1061.
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We then tested whether the effects of the mutations were additive. The double-mutated strains were constructed in the MC1061 background first by eliminating the cat gene as described in Materials and methods and then by transducing the second mutation. The elimination of the cat gene has no effect on the level of production of recombinant scFv17F12-PhoAGN (data not shown). Results obtained with the double-mutated strains are presented in Figure 3. No addition of the effects was observed when the wecB and wecE null mutations were combined. On the other hand, the effects of the wecB and wecE null mutations can be added to those of yheS, and we observed a 3- to 4-fold increase in production compared to the wild-type strain.
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The first example we tested was scFv17F12QUAD-PhoAGN, an engineered recombinant scFv17F12-PhoAGN that is produced more efficiently as a functional protein than the wild-type hybrid. As can be seen in Figure 5, the effects of the null mutations in yheS, wecB and wecE were not identical to those observed with the wild-type protein. Only mutations in wecB and wecE had a slight effect on the production (1020% up) and the yheS null mutation seemed to be ineffective in increasing functional antibody fragment production.
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The effects of yheS, wecB or wecE inactivation are additive to those of fkpA or skp overexpression
One interesting point was to examine if the effects of the mutations isolated in our study cumulate with those previously reported when fkpA or skp are in multicopy (Bothmann and Plückthun, 1998, 2000
). The fkpA and skp genes were expressed from their own promoter after cloning directly in the vector expressing the recombinant antibody fragment. Figure 7 shows that the presence of fkpA and skp in multicopy induced, respectively, 8- and 3.5-fold expression of functional scFv17F12-PhoAGN (compare dark grey and light grey bars in Figure 7). The addition of either null mutation had a positive effect. The best results were obtained with the yheS wecE double-null mutant when fkpA was in multicopy (
16-fold increase in functional scFv17F12-PhoAGN produced compared to MC1061; see Figure 7A) and with the yheS single mutant or yheS wecB double mutant when skp was in multicopy (
6-fold increase in functional scFv17F12-PhoAGN produced compared to wild-type MC1061; see Figure 7B).
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Discussion |
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First, this selection procedure has proved to be simple, rapid and efficient, since bacteria from the eight deep blue colonies selected from 10 000 clones obtained from a transposon insertions library actually produce higher amounts of functional recombinant antibody. For seven clones, it was shown that these higher productions are linked to the presence of the transposon. Secondly, this procedure appears to be sensitive enough since we isolated mutations that increase functional antibody fragment production 1.5- to 2-fold. This sensitivity is approximately the same as that described for phage display technology used for the selection of recombinant antibody with an engineered framework (Deng et al., 1994; Turner et al., 1997
; Tuckey and Noren, 2002
).
By using the above selection procedure, we looked for null mutations in the E.coli chromosome that would lead to an increase in functional recombinant scFv-PhoAGN. We isolated three different transposon insertions in yheS, wecB and wecE and showed that directed inactivation of each of these genes resulted in a 1.5- to 2-fold increase in the yield of periplasmic functional scFv17F12-PhoAGN. We demonstrated that the effects of null mutations in yheS could be cumulated with those occurring in either wecB or wecE and that each of these could also be positively combined with overexpression of skp or fkpA, two genes previously reported to be associated with positive effects on antibody production (Bothmann and Plückthun, 1998, 2000
). Thus, by combining null mutations and fkpA overexpression we obtained a 16-fold increase in production of functional recombinant antibody, an 8-fold increase being obtained with fkpA overexpression alone. We therefore demonstrated the possibility of engineering E.coli bacterial cells for the production of functional recombinant antibodies not only by overexpressing genes as already described but also by inactivating genes on the chromosome.
The modes of action of yheS on the one hand and wecB and wecE on the other hand appear different. First, as noted above, the effects of yheS mutation can be cumulated with those of wecB or wecE mutations. Secondly, the effects of the mutations on cellular distribution of different species of recombinant scFv-PhoAGN produced are not the same for yheS and wecB or wecE. To our knowledge, improving functional recombinant antibody production by any method is accompanied by an increase in the amount of total soluble recombinant antibody. This total soluble recombinant antibody fraction that is present in periplasmic extracts is composed of functional and non-functional recombinant scFv and increasing the amount of the total soluble antibody fraction results in an increase in the functional antibodies fraction (Kazemier et al., 1996; Boquet et al., 2000
). We observed that inactivation of wecB or wecE induces this type of response. This is not the case for the yheS null mutation which offers an original pattern of scFv-PhoAGN distribution in cellular extracts: while the amount of functional scFv-PhoAGN increases (ELISA results), the total soluble scFv-PhoAGN fraction remains unchanged independently of the fractionation method used (western blot analysis). At present, the reasons for the positive effects observed with yheS inactivation are unclear. This gene encodes a putative 637 residue cytoplasmic protein of unknown function that displays a 210 amino acid region of homology with the nucleotide-binding domain of ATP-binding cassette (ABC) transporter (NCBI CDD cd00267). It is part of an operon containing yheS yheT and yheU. We noted that yheT inactivation by cassette insertion has no effect on recombinant antibody production, suggesting that there is no polar effect with yheS inactivation (J.Dassa and P.Belin, unpublished results). However, this role appears quite peculiar, since mutation of this gene affects the production level of scFv17F12-PhoAGN but not of scFvM
2-3-PhoAGN.
The effects of wecB and wecE inactivation seem to be more general than for yheS inactivation. We demonstrated that the wecB and wecE null mutants are effective at increasing the production of another recombinant antibody scFvM2-3-PhoAGN. The wec gene cluster products are implicated in the synthesis of lipid III, a precursor of ECA, from undecaprenyl monophosphate. Lipid I and lipid II are intermediates of this reaction and wecB inactivation blocks the synthesis of lipid II from lipid I while wecE null mutation impedes synthesis of lipid III from lipid II (Meier-Dieter et al., 1990
). It is tempting, therefore, to anticipate that perturbation of the synthesis of ECA by null mutations might induce an increased production of functional recombinant antibody. This view is supported by the observation that the effects of the two mutations cannot be added. However, we noted that inactivation of wecA, whose product is implicated in the synthesis if lipid I from undecaprenyl monophosphate, does not result in an increase in functional recombinant antibody (data not shown). More precisely, the presence of the wecA mutation resulted in growth difficulties in different genetic backgrounds. Concerning the modes of action of wecE, it has been reported that wecE inactivation causes the accumulation of lipid II, and that accumulation of lipid II stimulates two extracytoplasmic stress response systems, the Cpx signal transduction pathway and the
E regulatory system (Danese et al., 1998
). Activation of these systems induces the expression of genes involved in the physiology of extracytoplasmic proteins, and notably fkpA and skp (Danese and Silhavy, 1997
; Dartigalongue et al., 2001
). Although we cannot exclude a possible effect of this activation on scFv-PhoAGN folding, several points remain unclear. How can we explain the similar mode of action of wecB and wecE null mutations on functional scFv17F12-PhoAGN production, while wecB inactivation leads to the accumulation of lipid I, a precursor of lipid II? Why is there a lack of effectiveness of the different mutations on functional scFv17F12(His)6 production? Further experiments with (i) null mutants blocked at other steps of ECA synthesis, (ii) null mutants of the Cpx pathway, (iii) strains containing reporter fusion of the Cpx signal transduction pathway or
E activity and (iv) different scFv(His)6 encoding plasmid constructions are currently under investigation to gather clues to the modes of action of yheS, wecB and wecE inactivation.
The characteristics of our selection procedure make it a method of choice for other applications. The first example appears as a continuation of our work and concerns E.coli cell engineering by selecting bacteria not carrying gene disruptions but chromosomal point mutations that would be advantageous for functional recombinant antibody production. During our work, we isolated the clone 2.3 which probably accumulated one or several mutations unlinked to the transposon and displayed better production of functional recombinant scFv. The recombinant plasmid isolated from clone 2.3 was as toxic to wild-type strain as the parental plasmid, suggesting that the mutation is not carried by the plasmid but more probably results from a chromosomal point mutation. The second example is the selection of recombinant antibodies with an engineered framework. To date, such antibodies have predominantly been selected by rational design (Knappik and Plückthun, 1995; Proba et al., 1998
; Wall and Plückthun, 1999
) or by phage display after mutagenesis of recombinant antibody cDNA (Deng et al., 1994
; Turner et al., 1997
; Tuckey and Noren, 2002
). However, selection and screening by phage display are often time-consuming, since these steps include several rounds of panning and ELISA with phage suspension obtained after growing individual colonies. With our screening procedure, several individual mutations could be isolated rapidly for their positive effects on functional recombinant antibody production. Finally, the possibility of using this screen with other proteins is attractive. Numerous proteins aggregate as inclusion bodies when overproduced and exhibit different degrees of toxicity (Kurland and Dong, 1996
; Baneyx, 1999
). By searching for toxicity reduction, we can envisage selecting (i) strains improved for functional protein production or (ii) protein variants evolved for efficient folding. This search for toxicity suppressors will imply the expression of the toxic protein fused with a partner displaying a detectable activity. In addition to PhoA that is restricted to periplasmic expression, several partners might be used to monitor cytoplasmic expression, like green fluorescent protein, ß-galactosidase or chloramphenicol acetyl transferase. Besides, these partners have been used fused to proteins as solubility reporters: the more the protein is insoluble, the less the reporter activity is detectable (Maxwell et al., 1999
; Waldo et al., 1999
; Wigley et al., 2001
). However, the use of such fusions for the screening of large libraries remains difficult in spite of several successes achieved by means of green fluorescent protein fusions (Waldo, 2003
). One of the most common problems with fusion proteins is the degradation occurring between the two partners of the fusion. In the case of solubility screening based on measurement of reporter activity, such degradation will lead to the delivery of a fully active soluble reporter protein, while the protein of interest will stay aggregated. Coupling such detection with a toxicity assay based on the diminution of the amount of aggregated protein in favour of folding should help to overcome these screening difficulties.
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Received May 5, 2004; accepted June 2, 2004.
Edited by Greg Winter