The E. coli strains produced in this work are derivatives of the commonly used expression strain BL21(DE3) (Studier and Moffatt, 1986). In addition to being widely utilized for expression, this strain also lacks the outer membrane protease OmpT, which we reasoned would be beneficial for over-expression of OMPs in particular. For transductions, we used the generally transducing bacteriophage P1 vir. The donor strains harboring the kanamycin cassettes for gene deletion were from the Keio collection (Baba et al., 2006). The K-12 reference strain was BW25113 (Datsenko and Wanner, 2000).

Bacteria were grown in lysogeny broth (LB) unless stated otherwise. For most experiments, we used the “Lennox” formulation: 10 g tryptone, 5 g yeast extract, and 5 g NaCl per liter. In the text below, we mean this formulation when referring to LB. For growth curves, we also used the “Miller” formulation (10 g tryptone, 5 g yeast extract, and 10 g NaCl per liter)—we refer to this as LB-Miller. Where necessary, media were supplemented with kanamycin (kan) or ampicillin (amp) at 25 μg/ml and 100 μg/ml, respectively. SOC medium was used after transformations as described elsewhere (Hanahan, 1983). As a defined medium, we used the minimal medium M9 (Miller, 1972) supplemented with glucose 0.2% (w/v) and 18 amino acids (all except cysteine and tyrosine) at 0.1 mg/ml each. Bacteria were usually grown at 37°C, unless harboring the plasmid pCP20, in which case they were grown either at 30°C for plasmid maintenance or 42°C to cure the plasmid. Some of the omp deletion strains, such as, the quadruple mutant BL21ΔABCF, grew significantly better at 30°C than at 37°C, and these were thus propagated at 30°C.

Plasmid pCP20 encoding the FLP recombinase (Flippase) was used for excising the kanamycin cassette after transduction into the knock-out strains (Cherepanov and Wackernagel, 1995). For λ red recombination, we employed the plasmid pKD46 (Datsenko and Wanner, 2000). The expression constructs used for testing our strains have been described previously: pET3b containing the genes encoding ompX and variants thereof (Arnold et al., 2007), pASK-IBA2 with Yersinia enterocolitica YadA membrane anchor domain (YadAM) (Wollmann et al., 2006) or Intimin (Oberhettinger et al., 2012). pET3b-OmpX was also modified for the purpose of this study by inserting a double haemagglutinin (HA)-tag with GSG linkers (GSGYPYDVPDYAGSGYPYDVPDYAGSG) in the position between S53 and S54 of OmpX for easier detection (pET3b-OmpX-HA). The insertion was created by site-directed mutagenesis (Byrappa et al., 1995) using the primers given in Table 1.

P1 phage transduction was performed essentially as described (Thomason et al., 2007). Briefly, the E. coli strains from the Keio collection were grown in LB medium to an optical density at 600 nm (OD₆₀₀) of 1.0 and infected with P1 vir phages at various dilutions (e.g., 10⁻⁵, 10⁻⁶, 10⁻⁷), mixed with 3 ml liquid top agar and poured onto pre-warmed LB plates. The following day, a semi-confluent plate was chosen and the top agar scraped off. This was mixed with 2 ml LB medium supplemented with a drop of chloroform, vortexed for 2 min and centrifuged for 10 min at 5,000 × g. Another drop of chloroform added to the supernatant, which was then stored at 4°C. The parent strain E. coli BL21 Gold (DE3) was infected with dilution series of each P1 phage lysate and the titer of the lysates was calculated from the number of plaques formed on the plates.

For the transduction experiments, E. coli BL21 Gold (DE3) and derivatives were grown in LB medium overnight, diluted 1:100 in the morning and grown until an OD₆₀₀ of ~1.0, and then supplemented with 10 mM CaCl₂ and mixed the P1 phage lysate harboring the kan cassette specific for the desired knock-out at a multiplicity of infection (MOI) of 0.5, assuming 10⁹ bacteria/ml culture. After incubating for 20 min at 37°C, the infection was stopped by addition of 100 mM sodium citrate pH 5.5. The mix was centrifuged for 2 min at 5,000 × g. The pellet was washed in LB medium supplemented with 100 mM sodium citrate and centrifuged as before. This washing step was repeated twice and then the bacteria were incubated for ~1 h at 37°C in LB containing 100 mM sodium citrate. Finally, the bacteria were centrifuged for 2 min at 4,000 × g diluted in 100 μl LB medium and streaked out on LB agar plates supplemented with kanamycin and 10 mM sodium citrate.

To delete the ompF locus, we amplified the FLP recognition target (FRT)-kanamycin cassette flanked by sequence directly outside the sequence coding for OmpF, using the Keio collection ΔompF strain and the same primer sequences described in that study (Baba et al., 2006). Insertion of the kan cassette was achieved by λ red recombination, essentially as described (Datsenko and Wanner, 2000): the plasmid pKD46 was transformed into recipient strains by electroporation, and transformants were selected for by plating on ampicillin and growing at 30°C. To introduce the kan cassette into the ompF locus, the pKD46-containing bacteria were grown to mid-log phase at 30°C, at which time the λ red genes were induced by the addition of 1 mM L-arabinose. After 1 h of induction, the cells were harvested and made electrocompetent. One hundred nanograms of PCR product was transformed into the cells by electroporation, after which cells were allowed to recover for 1 h at 30°C in SOC medium supplemented with 1 mM L-arabinose. Transformants were then selected for by growing on LB with kanamycin at 30°C. To remove pKD46, bacteria were grown on LB + kan at 37°C, and then tested for ampicillin sensitivity. Insertion of the kan cassette into the ompF locus was verified by colony PCR.

For excision of the kan cassette, kan-resistant transductants were transformed with the conditionally replicating plasmid pCP20 encoding the FLP recombinase by electroporation, following the procedure suggested by Baba et al. (2006). After electroporation the cells were quickly mixed with 1 ml SOC medium and incubated for 1 h at 30°C. The bacteria were then plated on LB agar plates with ampicillin and incubated overnight at 30°C. To cure pCP20, one amp-resistant colony was streaked out onto an LB agar plate and incubated at 42°C overnight. To screen for mutants strains, colonies were streaked out on LB plates containing kan, amp, and no antibiotic, respectively, using a grid, and then incubated at 37°C overnight. Clones sensitive to both kan and amp were chosen, and correct deletions were verified using colony PCR.

For verification of the right gene deletions in our mutants PCR with primers specific for the upstream and downstream region of the gene to be deleted was used. The primers sequences are given in Table 1. Colony PCR was performed using Taq polymerase (New England Biolabs) and 20 pmol of primer per reaction. A typical colony PCR program was as follows: initial denaturation for 3 min at 94°C, followed by 25 cycles of denaturation (30 s at 94°C), annealing (20 s 50°C), and extension [1.5 min (or 5 min with ompC common primers) at 70°C]. After a final extension of 5 min at 70°C, samples were mixed with loading buffer and applied to a 1% agarose gel (0.8% for ompC common primers). The primer pair used amplified the coding sequences of each locus are indicated in Table 1.

Outer membrane isolations were performed essentially as described in Leo et al. (2015). Briefly, 20 ml of an overnight culture at OD₆₀₀ 1.0 were pelleted and washed with 10 mM HEPES buffer at pH 7.4. To promote lysis, 0.1 mg/ml lysozyme was added, along with MgCl₂ and MnCl₂ to 10 mM and a pinch of DNase I (Sigma). The cells were then disrupted using a bead beater (SpeedMill Plus from Analytik Jena, Germany). The lysates were centrifuged at 15,600 × g for 30 s in a tabletop centrifuge and the supernatant was then moved to a fresh 2 ml microcentrifuge tube and centrifuged at 15,600 × g for another 30 min. The supernatant was decanted and the brownish membrane pellet resuspended in 400 μl 10 mM HEPES pH 7.4 with 1% (w/v) N-lauroyl sarcosine. The inner membranes were solubilized at room temperature (RT) for 30 min. Following this, the tubes were centrifuged at 15,600 × g for 30 min to pellet the outer membrane. The pellet was washed with 10 mM HEPES pH 7.4 and then resuspended in 30 μl HEPES buffer. For SDS-PAGE, 10 μl of 4 × non-reducing SDS sample buffer was added. Fifteen percentage SDS-PAGE gels were used for experimental verification of the knockout mutants at the protein level. For Coomassie G-250 (colloidal) staining of polyacrylamide gels, 12 μl of the outer membrane samples were used. For silver staining (Nesterenko et al., 1994), 8 μl of the outer membrane prep samples were run in a polyacrylamide gel and for Western blots 6 μl of the outer membrane prep samples were used.

To draw growth curves, starter cultures were grown in 5 ml LB or supplemented M9 medium and the OD₆₀₀ values were measured. The bacteria were then diluted to an OD₆₀₀ value of 0.01 and 5 μl of this suspension was added to 200 μl of medium (LB, LB-Miller, or supplemented M9) in a sterile microtiter plate. For blanks, no bacteria were added. The plates were sealed with a BreathEasy® membrane (from Sigma-Aldrich). The plates were incubated in a Biotek Synergy plate reader with controlled temperature and orbital rotation at the “slow” setting; absorbance at 600 nm was recorded at 20-min intervals. For plotting, values from four biological replicates were averaged.

For sedimentation assays, an overnight culture was diluted 1:100 in fresh LB (total volume 10 ml). The cultures were grown with shaking at 30°C in a flask till late log phase (OD₆₀₀ ~1.0), at which point MgCl₂ or CaCl₂ were added to 10 mM to some of the cultures. After a further hour of incubation at 30°C with shaking, the bacteria were transferred carefully to 18 mm tubes so as not to disrupt any floccules. The tubes were then incubated statically at RT. One hundred and fifty microliter samples were taken from the very top of the cultures at 5-min intervals for 20 min and the absorbance at 600 nm was measured using a microcuvette (light path 1 cm). To estimate autoaggregation, the absorbance at each time point was compared to the absorbance at time point zero and expressed as a percentage: (At ∗ 100)/A0, where A_t is the absorbance at the relevant time point and A₀ is the absorbance at time point zero. For plotting, three biological replicates were used.

For photography, 5 ml cultures were grown to late log phase (OD₆₀₀ ≈ 0.8) and MgCl₂ or CaCl₂ were added as above. After a further hour of shaking, the tubes were incubated statically at RT and photographed after 20 min and 2 h.

For inducing recombinant protein production, plasmids encoding the test proteins were transform into the quadruple mutant strain (BL21ΔABCF). For production, a 5 ml overnight culture of a transformed clone was diluted 1:100 in fresh LB and grown at 30°C till mid-log phase (OD₆₀₀ ~0.5). The culture was then induced with either isopropyl thiogalactoside (at 1 mM) or anhydrotetracycline (at 50 ng/ml). The cultures were incubated at 30°C for a further 2 h, after which the cells were harvested and outer membranes were isolated. The OMPs were separated by SDS-PAGE. Over-expressed YadAM, OmpX, and its duplicated variant OmpX88 were detected by colloidal Coomassie G-250 staining. Overexpression of a HA-tagged variant of OmpX and Intimin was detected by immunoblotting. The proteins were transferred to a 0.45 μm polyvinylidine difluoride membrane (Thermo Scientific) using a semi-dry transfer unit (Hoefer TE70X). The membranes were blocked with 2% skimmed milk powder dissolved in phosphate-buffered saline (PBS; 20 mM sodium phosphate pH 7.4, 150 mMNaCl) for 1 h at RT or overnight at 4°C. This was followed by incubation for an hour with primary antibodies; 1:2,000 dilution of rabbit anti-HA tag antibody (for OmpX-HA) and 1:5000 dilution of rabbit anti-Intimin antibody (Oberhettinger et al., 2012). The membrane was washed three times with PBS+0.05% Tween20 followed by incubation with a 1:10,000 dilution of goat anti-rabbit IgG horseradish peroxidase-conjugate (Santa Cruz Biotech) in PBS+2% skimmed milk powder for an hour. The membrane was washed three times with PBS+0.05% Tween20. The bands were detected using enhanced chemiluminescence (Pierce ECL western blotting substrate) and a Kodak 4000R Image station.

For quantitative examination of Intimin and OmpX-HA expression, we performed whole-cell enzyme-linked immunosorbent assays (ELISAs). Bacteria transformed with the corresponding plasmids were grown overnight at 30°C in 5 ml LB medium with ampicillin. The following day, the bacteria were diluted 1:10 in fresh medium (5 ml, with amp) and grown till mid-log (OD₆₀₀ ~0.5; about 2 h), at which time they were induced with anhydrotetracycline or isopropyl thiogalactoside as in the section on Recombinant Protein Expression and Detection. The bacteria were grown at 30°C for another 2 h. The OD₆₀₀ of the cultures was measured and the bacteria were diluted in PBS to an OD₆₀₀ value of 0.2. One hundred microliters of this suspension were applied to the wells of a polystyrene microtiter plate and bacteria were allowed to adhere to the surface of the well for 1 h at RT. The wells were washed three times with 200 μl washing buffer [PBS + 0.1% bovine serum albumin (BSA, from VWR)] and then blocked for 1 h with PBS + 2% BSA. The wells were washed once with 200 μl washing buffer, and 100 μl of the primary antibody diluted into blocking buffer was applied. The antibodies were the same as in the section on Recombinant Protein Expression and Detection, anti-HA (1:2,000), and anti-Intimin (1:1,000). After an hour's incubation, the wells were washed three times as above, and the secondary antibody (anti-rabbit-HRP, from Agrisera) was added, diluted 1:2,000 in blocking buffer. The plate was incubated for 1 h at RT, after which the wells were washed three times as above. Detection was performed using the colorimetric HRP substrate ABTS (ThermoScientific) according to the manufacturer's instructions. After color development (40 min) the reactions were stopped with 1% SDS and absorbances were recorded at 405 nm.

To avoid the problems we had encountered with the strains from Prilipov et al. we pursued a different strategy in making our knock-out strains. We decided to delete the entire coding sequences for the four genes encoding the OMPs lacking in the Omp8 strain of Prilipov et al. namely ompA, ompC, ompF, and lamB (Prilipov et al., 1998b). We reasoned that this strategy would prevent the possibility of a genetic reversion, thus improving the genetic stability of the newly generated knock-out strains. BL21 strains probably do not express ompC naturally, due to an insertion element-mediated deletion of the upstream region of the ompC locus, including the signal peptide of OmpC (Pugsley and Rosenbusch, 1983; Studier et al., 2009; Han et al., 2012). Nevertheless, we decided to delete the entire ompC coding sequence to fully prevent the possibility of reversion, e.g., by a recombination event restoring a functional signal peptide. To make the deletions, we employed the Keio collection, a set of 3,985 single-gene deletions in E. coli, where virtually the entire coding sequence of the deleted genes is replaced by a kanamycin resistance cassette (Baba et al., 2006). This cassette is flanked by sequences targeted by the FLP recombinase; thus, when FLP is supplied in trans, the kan cassette can be excised from the genome leaving only a ~100 bp-long scar sequence.

We produced the single knock-outs in E. coli BL21 Gold (DE3) by phage P1 transduction: a phage lysate was produced from the Keio strains harboring the desired OMP gene deletions. This lysate was used to infect the recipient strain, and kan-resistant transductants were then selected for by plating on kanamycin plates. To remove the kan cassette, we introduced the FLP-containing plasmid pCP20 into the kan-resistant bacteria and selected for amp-resistant colonies without the addition of kanamycin. To cure pCP20, clones were plated onto LB (no selection) and grown at 42°C. Clones that were sensitive to both amp and kan were chosen for PCR screening to verify the loss of the kan cassette and the loss of the wild-type allele. Using this strategy, we produced the four individual knock-outs, and then by re-iterating the process, we obtained five double knock-outs, three triple knock-outs, and the quadruple knock-out strain (Table 2).

We experienced particular problems producing the ompA-ompF double knock-out and the ompA-ompF-ompC triple knock-out. The ompA and ompF loci are situated relatively close to each other in the E. coli genome (distance ~32 kb; P1 can transduce fragments ~ three times this size); therefore, we often obtained revertants for one mutant while trying to produce the other. To circumvent this problem, we employed a different strategy: we amplified the FRT-kan cassette from the Keio ΔompF strain by PCR, with 50 bp overhangs identical to the sequence flanking the ompF coding region, as was used to make the ompF mutant in the original Keio collection (Baba et al., 2006). We then produced the BL21ΔAF and BL21ΔACF strains by λ red recombination in the BL21ΔA and BL21ΔAC strains (Datsenko and Wanner, 2000). We then selected for kan-resistant colonies and subsequently removed the kan cassette as described for the transduction experiments. Indeed, this strategy proved successful.

To verify that the correct gene had been deleted, we amplified the target coding sequence by colony PCR. Using primers specific for each of the four loci, we could show that the mutant strains had lost the wild-type allele (Figure 1A). The sequences remaining in the deletion mutants correspond to ~150 bp, the size of the scar sequence and flanking regions, indicating that the gene was actually replaced by the kan cassette, which in turn was excised by FLP. The size of the wild-type coding sequence is 1041 bp for ompA, 1089 bp for ompF, 1104 bp for ompC, and 1341 bp for lamB. These sizes are consistent with the PCR products obtained from the parent strain BL21 Gold (DE3). For verification of the ompC deletion, another forward primer binding downstream the rcsC locus was used due to the fact that in E. coli BL21 the upstream region of ompC is deleted (Studier et al., 2009). Thus, the forward primer amplifying the ompC locus in E. coli K-12 strains does not produce a product in B strains. The common forward primer, which binds downstream of the rscC gene, produces a product in both strains when combined with the same reverse primer used for amplifying the K-12 locus (Table 1). In BL21 Gold (DE3), this results in a product of ~1.9 kb. In K-12, the product also contains the region deleted in BL21 Gold (DE3), so the total length of the product is ~5.3 kb. For the mutants, where the ompC coding sequence is deleted within the K-12 context, the product is ~4.2 kb (Figure 1B). These results confirm that the kan cassette had correctly replaced the ompC sequence of BL21 Gold (DE3).

In order to show that the PCR-positive knock-out strains lack the corresponding proteins, we prepared outer membrane samples from all the strains and analyzed these by SDS-PAGE and silver staining (Figure 1C). E. coli BL21(DE3) was included as a positive sample showing outer membrane protein bands of interest. Maltoporin (LamB) has a size of ~49 kDa. OmpC and OmpF are approximately the same size and run as a single band at ~39 kDa, and OmpA has the smallest size of ~35 kDa. All of these protein bands are visible in the parent strain. The single knock-out strain BL21ΔA shows all bands of interest except the 35 kDa OmpA band indicating that this protein is lacking in this strain. The ΔB strain shows the bands of interest at 35 kDa for OmpA and at 37 kDa for OmpC and OmpF but no band at 49 kDa. As a note, maltoporin is not well expressed in E. coli B strains, including BL21, which makes the confirmation of the lamB knock-out difficult on the protein level (Ronen and Raanan-Ashkenazi, 1971; Han et al., 2012); thus, the lamB knock-outs could only be fully confirmed by PCR. Single ompC and ompF knock-outs show a reduction in band intensity at 39 kDa, and only the double knock-outs lack the band at 39 kDa completely, suggesting that our strategy of knocking out ompC was a reasonable approach to make sure the gene is entirely inactivated. A possible explanation for a residual band at 39 kDa (e.g. for BL21ΔF) is expression of the cryptic porin OmpN (Prilipov et al., 1998a). Taken together, the PCR results and OMP profiles of the strains show that we have successfully deleted the genes encoding the major OMPs, either singly or in all combinations.

While working with the new strains, it became obvious that by altering the outer membrane protein composition the growth behavior changed. BL21ΔABCF grew significantly more slowly than the parent strain BL21(DE3). The strain grew initially faster at 37°C than at 30°C, but the culture at 37°C saturated at a lower OD₆₀₀ (Figure 2A). However, even at 30°C, BL21ΔABCF did not reach the same OD₆₀₀ value as BL21(DE3).

We also observed that when using the Miller formulation of LB (with 10 g sodium chloride per liter), BL21ΔABCF grew significantly slower than in our standard LB (5 g NaCl/l), also at 30°C, whereas BL21(DE3) showed no significant differences in growth in the two media (Figure 2B). We do not currently have an explanation for this phenomenon; perhaps BL21ΔABCF is unable to compensate efficiently for the increased osmolarity of LB-Miller.

We also tested the growth of BL21ΔABCF in defined medium (Figure 2C). For this, we used minimal medium M9 supplemented with 18 amino acids. Though growth was slower than in LB, BL21ΔABCF reached higher optical densities than in the rich, undefined medium. It apparently also reached a higher density than BL21(DE3); however, part of the higher absorbance readings of the quadruple mutant cultures could be attributed to the tendency of BL21ΔABCF to clump in the defined medium, presumably due to the relatively high concentration of magnesium (2 mM; see section Aggregation of BL21ΔABCF in the Presence of Divalent Cations).

As noted above, BL21ΔABCF tends to aggregate in the presence of divalent cations. When Mg²⁺ or Ca²⁺ is added to the medium, BL21ΔABCF flocculates and settles rapidly at the bottom of the tube under static condition (Figures 3A,B). In contrast, the parent strain does not aggregate in the presence of either ion during the short time frame of the experiment (Figures 3A,C). However, upon prolonged incubation (>1 h), also BL21(DE3) began to flocculate in the presence of Ca²⁺ (Figure 3D). The reason for the rapid aggregation of BL21ΔABCF is not clear. A possibility might be the increased binding of divalent cations by the more exposed lipid A phosphates in the BL21ΔABCF strain, leading to electrostatic attraction through bridging divalent cations. In principle, the aggregation caused by CaCl₂ might cause problems when preparing chemically competent cells; however, in our hands BL21ΔABCF can be made competent and transformed efficiently using the standard CaCl₂ protocol (data not shown).

To qualitatively test the performance of our new quadruple mutant strain, we over-expressed four test proteins from our laboratory. For comparison, we used the parent strain E. coli BL21 (DE3). The test proteins were OmpX, a native OMP of E. coli, a duplicated variant of this protein with 16 β-strands rather than the usual eight (Arnold et al., 2007), and another OmpX variant containing a HA tag in one of the extracellular loops. Additionally, we tested the expression of two autotransporter proteins: the membrane anchor domain of the Yersinia adhesin YadA (Wollmann et al., 2006) and the inverse autotransporter Intimin from enteropathogenic E. coli (Oberhettinger et al., 2012).

The expression of wild-type OmpX can be seen as a band at 15 kDa when stained with Coomassie G-250 (Figure 4A). The band is more intense for BL21ΔABCF than for the parent strain. Other bands of native proteins, for example the band at 18 kDa that presumably represents OmpW, show that the same amount of sample was loaded. Note that there is no clear difference between the strains for the duplicated variant OmpX88 (~37 kDa). The OmpX-HA construct was also expressed in BL21(DE3) and BL21ΔABCF. A Western blot of outer membrane preparations (Figure 4B) shows that for the quadruple knock-out strain, the total amount of expressed OmpX-HA is higher compared to the parent strain BL21(DE3). An additional band was detected at an increased molecular weight (~25 kDa) that we originally attributed to non-denatured OmpX-HA (Figure 4B, right panel), similar to previous findings on OmpX gel shifts (Arnold et al., 2007), Based on the OMP gel shift phenomenon (Rosenbusch, 1974; Schweizer et al., 1978), we know that native and denatured forms of OMPs can migrate differently in SDS-PAGE. However, when comparing heated and unheated samples, the band did not change, suggesting that it is an artifact yet to be explained (data not shown). The third specific band at ~50 kDa is presumably a folded dimer of OmpX at very low concentration (Chaturvedi and Mahalakshmi, 2013).

YadA is an obligate homotrimer and an adhesin of enteropathogenic Yersiniae (Mühlenkamp et al., 2015). It is an extremely stable protein which remains trimeric in the presence of denaturants such as, SDS and urea (Wollmann et al., 2006). In SDS-PAGE, YadAM (membrane anchor domain of YadA) migrates at an apparent molecular weight of 45 kDa (Wollmann et al., 2006). The expression of YadAM in BL21(DE3) and BL21ΔABCF is shown in Figure 4C. The colloidal Coomassie G250-stained gel shows better expression of YadAM in BL21ΔABCF than BL21(DE3). As mentioned above, many native outer membrane protein bands (e.g., ~30 kDa and ~60 kDa) are absent in the ΔABCF strain.

The expression of Intimin is shown in Figure 4D. Here, a construct including a StrepII tag was used and its expression visualized specifically in a Western blot using an anti-Intimin antibody (Oberhettinger et al., 2012). The blot shows two bands for the parent strain: one at ~95 kDa corresponding to the molecular weight of Intimin and a second band at ~120 kDa. This latter band is sometimes observed in Intimin blots, though its provenance is not clear (Heinz et al., 2016; Leo et al., 2016). For the ΔABCF strain, the 95 kDa band is more intense, and the ~120 kDa band fainter, suggesting better membrane insertion. In addition, some apparent degradation product can be seen for the quadruple mutant (band at ~70 kDa). The superiority of the BL21ΔABCF strain is demonstrated by a higher yield of the “correct” Intimin band and less of the (presumably mis-incorporated) 120 kDa band.

To gain a more quantitative view of OMP production in BL21ΔABCF and to assess reproducibility between culture batches, we performed whole-cell ELISA on bacteria expressing OmpX-HA and Intimin. We compared expression of these two proteins in BL21(DE3), BL21ΔABCF, and the Prilipov strain BL21 Omp8. We also tested the expression of YadAM in these strains, but due to technical problems with detecting the StrepII tag on the bacterial surface combined with the tendency of YadAM-expressing cells to clump, we did not obtain reliable results (data not shown). In this construct, only a short stretch of YadAM is exposed to the surface, so the StrepII tag is presumably not fully accessible to antibodies (Shahid et al., 2012).

BL21ΔABCF produced both Intimin and OmpX-HA at higher amounts than the parent strain BL21(DE3) (Figure 5). Somewhat unexpectedly, BL21ΔABCF also outperformed the Omp8 strain, though the difference is not large. The ELISA also demonstrate that the variability between biological replicates is low, though for Intimin there is more variability between replicates. Incidentally, in the whole-cell assays the detected proteins must be surface-exposed, showing that the detected species must be correctly processed and inserted into the membrane.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. This work has been included in a patent application (U.S. Patent Office application number 62438094).