Eur Biophys J (2015) 44:503–512 DOI 10.1007/s00249-015-1032-y

ORIGINAL PAPER

Modulating bilayer mechanical properties to promote the coupled folding and insertion of an integral membrane protein Michaela Herrmann1 · Bartholomäus Danielczak1 · Martin Textor1 · Jessica Klement1 · Sandro Keller1 

Received: 11 March 2015 / Revised: 23 April 2015 / Accepted: 5 May 2015 / Published online: 29 May 2015 © European Biophysical Societies’ Association 2015

Abstract  Bilayer mechanical properties are not only of crucial importance to the mechanism of action of mechanosensation in lipid membranes but also affect preparative laboratory tasks such as membrane-protein refolding. We report this for coupled refolding and bilayer insertion of outer membrane phospholipase A (OmpLA), an integral membrane enzyme that catalyses the hydrolytic cleavage of glycerophospholipids. OmpLA can be refolded into a variety of detergent micelles and unilamellar vesicles composed of short-chain phospholipids but, in the absence of chemical or molecular chaperones, not into thicker membranes. Controlled modulation of bilayer mechanical properties by judicious use of subsolubilising concentrations of detergents induces monolayer curvature strain, acyl chain fluidisation, membrane thinning, and transient aqueous bilayer defects. This enables quantitative and functional refolding of OmpLA even into bilayer membranes composed of long-chain phospholipids to yield enzymatically active proteoliposomes without requiring membrane solubilisation. Keywords  Gel-shift assay · Membrane enzymes · Outer membrane phospholipase A · Proteoliposomes · Protein refolding

Special issue: Biophysics of Mechanotransduction. * Sandro Keller [email protected] 1



Molecular Biophysics, University of Kaiserslautern, Erwin‑Schrödinger‑Str. 13, 67663 Kaiserslautern, Germany

Introduction Cells have sophisticated mechanisms enabling sensing and response to changes in the bilayer mechanical properties of their membranes (Kung et al. 2010; Anishkin et al. 2014). For instance, when the cytoplasm of a bacterial cell turns hyperosmotic with regard to the extracellular medium, water influx results in swelling of the cell and stretching of the cell membrane. This mechanical strain leads to lateral expansion and concomitant thinning of the bilayer membrane. At the molecular level, these two coupled processes are accomplished by fluidisation of lipid acyl chains, which assume a more disordered, dynamic conformation occupying more membrane area but having a reduced effective chain length. By simultaneously undergoing lateral expansion and contraction along its normal, the lipid bilayer thus partly relaxes in-plane mechanical stress without a major volume change, which would incur a much higher freeenergy penalty. However, the membrane’s ability to cope with mechanical stress is limited, and proteins such as the mechanosensitive channel of large conductance (MscL) from Escherichia coli therefore act as emergency valves to enable release of osmolytes before further osmomechanical stress would cause membrane rupture and cell lysis (Perozo and Rees 2003; Martinac et al. 2014). Seminal experiments (Sukharev et al. 1993; Perozo et al. 2002; Sukharev 2002; Moe and Blount 2005) on mechanosensitive channels reconstituted into artificial lipid membranes have demonstrated that it is the changes in bilayer mechanical properties themselves that trigger the conformational rearrangements leading to channel opening. For MscL, it has been shown (Perozo et al. 2002) that opening is caused primarily by alterations in the transbilayer lateral pressure profile induced by strain, whereas hydrophobic mismatch due to bilayer thinning seems to be of secondary

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importance only. In vitro, opening of this and other mechanosensitive channels can be triggered by asymmetric incorporation of subsolubilising concentrations of lysophosphatidylcholines (LPCs) into one bilayer leaflet (Lundbaek and Andersen 1994). This not only provides a convenient experimental technique for studying mechanosensation by use of well-defined proteoliposomes but also furnishes insights into the nature of the bilayer-deformation forces responsible for channel opening, although the underlying mechanisms seem to be different (Mukherjee et al. 2014). It may seem surprising that the effects of mechanical stress within the bilayer plane, which ultimately originate from osmotic pressure, are partially emulated by detergents such as LPCs. In this context, it should be realised that, even well below the concentrations required for the much more drastic process of membrane solubilisation, detergents have a profound effect on bilayer mechanical properties. This is manifested in a multitude of interrelated phenomena, including monolayer and, for asymmetric detergent insertion, bilayer curvature strain, acyl chain disordering, lateral expansion, and bilayer thinning, which give rise to accelerated transbilayer lipid movement (so-called flip–flop), increased permeability to polar solutes and ions, membrane (hemi)fusion, and vesicle agglomeration (Pantaler et al. 2000; Heerklotz 2008; Ahyayauch et al. 2010; Lichtenberg et al. 2013). Unravelling and disentangling these processes is crucial not only for a better understanding of mechanosensation but also for controlled application of bilayer deformation forces in preparative applications requiring judicious modulation of bilayer mechanical properties. Herein, we report one such example, namely, the coupled in-vitro refolding and membrane insertion of an integral membrane protein. In the course of this process, polar and charged protein moieties must be translocated across the low-dielectric hydrocarbon core of the lipid bilayer to enable the protein to adopt its native transmembrane topology. The approach is exemplified for the functional refolding of E. coli outer membrane phospholipase A (OmpLA), an integral membrane enzyme that, upon activation, catalyses the hydrolytic cleavage of acyl chains in both the sn-1 and sn-2 positions of glycerophospholipids (Scandella and Kornberg 1971; Doi et al. 1972). OmpLA is a robust and well-studied model system for both experimental (Moon and Fleming 2011; Moon et al. 2011) and computational (Fleming et al. 2012) protein-folding studies, not least because it can be refolded from its urea-unfolded state into detergent micelles (Dekker et al. 1995) or into lipid bilayers made up of short-chain phospholipids (Burgess et al. 2008; Moon and Fleming 2011; Moon et al. 2011; Gessmann et al. 2014). By contrast, direct refolding into membranes composed of lipids with acyl chains longer than 12 carbon atoms has remained unsuccessful (Burgess et al. 2008). We

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show that modulation of bilayer mechanical properties by use of subsolubilising detergent concentrations transiently compromises the barrier function of the membrane, thus enabling quantitative refolding of OmpLA into preformed long-chain phospholipid bilayer vesicles to produce enzymatically active proteoliposomes.

Materials and methods Materials All chemicals were purchased in the highest available purity. CaCl2, 1,4-dithiothreitol (DTT), ethylenediamineN,N,N′,N′-tetraacetic acid (EDTA), glycine, HCl, isopropyl-β-d-thiogalactopyranoside (IPTG), MgCl2, and tris(hydroxymethyl)aminomethane (Tris) were from Carl Roth (Karlsruhe, Germany), Benzonase and sucrose were from Merck (Darmstadt, Germany), and poly(ethylene oxide) dodecyl ether (Brij-35), 5,5′-dithio-bis-(2-nitrobenzoic acid) (DTNB), and lauryldimethylamine N-oxide (LDAO) were from Sigma–Aldrich (Steinheim, Germany). Urea was obtained from Affymetrix (Santa Clara, USA), 2-hexadecanoylthio-1-ethylphosphorylcholine (HEPC) from Biozol (Eching, Germany), and N-dodecyl-N,Ndimethyl-3-ammonio-1-propanesulphonate (SB3-12) from Anatrace (Maumee, USA). n-Dodecylphosphocholine (DPC) and all n-alkyl-β-d-glucopyranosides and n-alkylβ-d-maltopyranosides were purchased from Glycon Biochemicals (Luckenwalde, Germany). 1,2-Dilauroyl-snglycero-3-phosphocholine (DLPC) was from Avanti Polar Lipids (Alabaster, USA) and 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) from Lipoid (Ludwigshafen, Germany). Protein production and purification The pldA gene encoding OmpLA without signal sequence was amplified from the E.  coli genome by polymerase chain reaction and cloned into a pET-24a(+) expression vector (Merck), as confirmed by bidirectional sequencing (SEQ-IT, Kaiserslautern, Germany) using T7 promoter and terminator primers. Lysogeny broth medium was inoculated with transformed E.  coli BL21(DE3) cells before production of inclusion bodies containing tag-free OmpLA was induced by addition of 0.4 mM IPTG with agitation at 37 °C for 4 h. Cells were harvested, washed, weighed, and stored at −80 °C. Inclusion bodies were isolated by thawing cell pellets on ice and resuspending them in 10 mL/(g wet biomass) ice-cold breakage buffer (50 mM Tris, 2 mM MgCl2, 40 mM EDTA, 25 % (w/v) sucrose, 0.01 % (v/v) Benzonase, pH 8.0). Cells were sonicated in an S-250A sonifier (Branson Ultrasonics, Danbury, USA) using two

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runs of 10 min each and, after addition of 0.01 % (w/v) Brij-35, a final run of 1 min at an amplitude of 40 %. The lysate was centrifuged for 45 min at 4 °C and 4500 g, and the pellet was resuspended in 40 mL washing buffer (10 mM Tris, 1 mM EDTA, pH 8.0), centrifuged for 30 min at 4 °C and 4500 g, and dissolved in solubilisation buffer (20 mM Tris, 2 mM EDTA, 8 M urea, 100 mM glycine, pH 8.3) with agitation for 3 h at 4 °C. Unfolded OmpLA was separated from residual impurities by centrifugation for 30 min at 4 °C and 4500 g. The protein concentration in the supernatant was ~3 mg/mL, as determined spectrophotometrically by using a molar extinction coefficient of 82.3/ (mM cm) at 280 nm (http://expasy.org/tools/protparam. html). Aliquots (2 mL) of the supernatant were shock-frozen in liquid nitrogen and stored at −80 °C.

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urea, 10.8 mM glycine, and appropriate concentrations of detergent or lipid and detergent (discussed in the section “Results and discussion”). The refolding mixture was further agitated for 16 h at 50 °C and 750 rpm on a Thermomixer compact (Eppendorf, Hamburg, Germany). Refolded protein and unfolded or misfolded protein were separated by SDS polyacrylamide gel electrophoresis (SDS-PAGE) by use of a NuPAGE Bis–Tris system (Life Technologies, Carlsbad, USA) with a PA gradient of 4–12 %. To this end, 1 µg reduced protein was applied to each slot before separation was performed at a constant voltage of 200 V. Gels were photographed with a C4000Z camera (Olympus, Tokyo, Japan), and protein bands were quantified densitometrically with ImageJ (Schneider et al. 2012) to determine the relative amount of refolded protein as a percentage of total protein.

Vesicle preparation and characterisation Enzyme activity assay Large unilamellar vesicles (LUVs) composed of DLPC or POPC were produced by 35-fold extrusion through two stacked polycarbonate filters with a pore diameter of 100 nm by use of a LiposoFast extruder (Avestin, Ottawa, Canada). Extruded vesicles had a unimodal size distribution with a mean diameter and standard deviation of (140 ± 10) nm for DLPC and (160 ± 10) nm for POPC, as derived from intensity-weighted size distributions obtained by dynamic light scattering (DLS) with a Zetasizer Nano S90 (Malvern Instruments, Worcestershire, UK). Small unilamellar vesicles (SUVs) composed of DLPC were produced by indirect sonication by use of a beaker resonator (Bandelin electronic, Berlin, Germany), as described in detail elsewhere (Klingler et al. 2015). Briefly, a flat-bottomed 5-mL glass vial containing 500 µL multilamellar vesicles was placed in the water-filled beaker resonator, which was thermostatted to 20 °C. The lipid suspension was sonicated twice for 15 min and twice for 10 min at a sonication power of 100 % in a sequence of 10-s pulses interrupted by 1-s pauses (Klingler et al. 2015). Intensityweighted size distributions revealed a dominant peak that accounted for >90 % of the scattering intensity with a mean diameter and standard deviation of (35 ± 1) nm. SUVs were stored at room temperature and used within one day.

Ca2+-triggered enzymatic activity was assessed by following the kinetics of OmpLA-catalysed hydrolysis of HEPC, an acylthioester analogue of LPCs (Aarsman et al. 1976). Hydrolysis releases thiol groups, which rapidly and stoichiometrically react with DTNB to form 2-nitro-5-thiobenzoate with a high molar extinction coefficient of 13.6/(mM cm) at 412 nm (Ellman 1959). Refolded OmpLA at a protein concentration of 4–33 µg/ mL was incubated with 1 mM HEPC and 0.8 mM DTNB for 2 h at room temperature in the dark before enzymatic hydrolysis of HEPC was initiated by addition of CaCl2 to reach final concentrations of 3.9–32.4 µg/mL OmpLA, 0.98 mM HEPC, 0.78 mM DTNB, and 19.6 mM CaCl2. Absorbance at 412 nm was monitored at room temperature by use of a V-630 UV–visible spectrophotometer (Jasco, Groß-Umstadt, Germany) with a 3-mm quartz glass cuvette (Hellma, Müllheim, Germany), and the normalised specific activity was determined as the time derivative of absorbance averaged over the first 250 s divided by the above molar extinction coefficient, optical pathlength, and concentration of refolded OmpLA. Within the range tested, the normalised specific activity was independent of OmpLA concentration.

Protein refolding into micelles and bilayer vesicles

Isothermal titration calorimetry

Detergent micelles or unilamellar vesicles consisting of both lipid (i.e., DLPC or POPC) and detergent (i.e., LDAO or OG) at different subsolubilising detergent concentrations were incubated in refolding buffer (20 mM Tris, 2 mM EDTA, pH 8.3) for 10 min at 50 °C to enable equilibration. When not indicated otherwise, unfolded OmpLA was refolded by drop dilution under agitation at 50 °C to reach final concentrations of 0.33 mg/mL OmpLA, 0.87 M

A DLPC/LDAO phase diagram at 25 °C and full hydration was derived from solubilisation and reconstitution experiments monitored by isothermal titration calorimetry (ITC), as described in detail elsewhere (Heerklotz et al. 2009; Textor et al. 2015). For solubilisation, 40, 100, or 60 mM  LDAO was titrated into suspensions of, respectively, 4, 6, or 8 mM  DLPC in the form of LUVs. For reconstitution, 20, 30, or 40 mM DPLC LUVs were titrated into 3,

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Fig. 1  Refolding yield and normalised specific activity of OmpLA in different detergents. a Fractions of OmpLA refolded at 50 °C into 27.3 mM OG, 7.8 mM NG, 26.8 mM OM, 9.3 mM NM, 3.5 mM DM, 1.7 mM UM, 1.2 mM DDM, 2.9 mM LDAO, 2.8 mM DPC, or 5.2 mM SB3-12. Refolding yields were obtained by densitometric

quantification of polyacrylamide gels. b Normalised specific activities at room temperature of the same samples containing refolded OmpLA. Error bars indicate standard deviations from 3 or, for LDAO, 20 independent refolding trials

4, or 5 mM LDAO, respectively. All experiments were performed on an iTC200 (Malvern Instruments) using a reference power of 25 µJ/s, a stirrer speed of 1000 rpm, a filter period of 5 s, an injection volume of 0.8 µL, and time spacings of 180 s. For analysis of raw thermograms, automated baseline assignment by singular value decomposition and peak integration were performed with NITPIC (Keller et al. 2012). To establish the phase diagram, the LDAO concentrations at the inflection points of both solubilisation and reconstitution isotherms were plotted against the corresponding DLPC concentrations, and the saturation (SAT) and solubilisation (SOL) boundaries were obtained from global linear fits to these data, as described elsewhere (Heerklotz et al. 2009; Textor et al. 2015).

included n-octyl-β-d-glucopyranoside (OG), n-nonyl-β-dglucopyranoside (NG), and a homologous series of n-alkylβ-d-maltopyranosides bearing octyl (OM), nonyl (NM), decyl (DM), undecyl (UM), or dodecyl (DDM) chains. Headgroup effects were investigated by comparing a series of dodecyl chain detergents including “harsh”, anionic SDS and zwitterionic LDAO, DPC, and SB3-12. For each detergent, ~3 mg/mL (~100 µM) unfolded OmpLA solubilised in 8 M urea was diluted at 50 °C with refolding buffer (20 mM Tris, 2 mM EDTA, pH 8.3) containing detergent at a concentration ~1 mM above the respective critical micellar concentration (CMC) determined in the presence of 1 M urea (Broecker and Keller 2013; Broecker et al. 2014). This ensured comparable concentrations of micellar detergent irrespective of the vastly diverging CMC values of the detergents used and accounted for the fact that the CMC depends on both temperature and urea concentration (Broecker and Keller 2013; Broecker et al. 2014). The choice of temperature was motivated by an initial refolding screen, which revealed that the optimum was 50 °C (data not shown, but see below for refolding into bilayers). Final protein and urea concentrations were 0.33 mg/mL (10.7 µM) and 0.87 M, respectively. The fraction of refolded protein was then quantified with a gel-shift assay (Inouye and Yee 1973; Nakamura and Mizushima 1976) based on the faster-than-expected migration of the natively folded protein in cold SDS-PAGE. Refolding yields thus obtained varied substantially, ranging from 0 for NG and SDS to >50 % for SB3-12 (Fig. 1a). Within the alkyl maltoside series, the yield amounted to ~40 % for NM and decreased for both shorter and longer alkyl chains. A different picture of detergent preference emerged on determination of the specific enzymatic activity, that is, the activity normalised relative to the concentration of refolded OmpLA. To this end, we used a colorimetric assay (Aarsman et al. 1976), in which we monitored the hydrolysis of

Results and discussion Refolding and activity in detergent micelles Similarly to other outer membrane proteins (Burgess et al. 2008; Dewald et al. 2011), OmpLA is soluble as a largely unfolded polypeptide chain in aqueous solutions of the chemical denaturants urea and guanidinium chloride. This has been exploited for both analytical (Moon and Fleming 2011) and preparative applications; the most prominent examples of the latter include solubilisation of recombinantly produced inclusion bodies and functional refolding of OmpLA into detergent micelles (Dekker et al. 1995, 1997) or short-chain phospholipid vesicles (Burgess et al. 2008). To complement earlier work (Dekker et al. 1995, 1997) and establish a reference framework for refolding experiments involving bilayer vesicles (see below), we first screened a set of popular detergents for their ability to support the functional refolding of OmpLA. Detergents with sugar-based headgroups

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the artificial substrate HEPC and the reaction of its product with DTNB to form a yellow-coloured complex. The alkyl maltoside detergents gave rise to low activities of 20 µmol/(min mg) under all conditions tested (data not shown, but see below for refolding into thicker bilayers), attesting to the success of the refolding procedure in generating enzymatically active proteoliposomes. Because temperature has been reported to have a strong effect on the bilayer refolding yield of OmpLA (Burgess et al. 2008) and other outer membrane proteins (Burgess et al. 2008; Maurya et al. 2013), we also performed a temperature screen (Fig. 3c). Raising the temperature from 7 to 50 °C increasingly facilitated OmpLA association with and insertion into LDAO-doped DLPC membranes, but this effect seemed to be counteracted by deteriorating protein stability at higher temperatures, as indicated by a reduced refolding yield at 60 °C. Thus, the temperature optimum for refolding into LDAO-containing DLPC bilayers was ~50 °C, as was observed for micellar refolding.

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Effect of free detergent and protein on bilayer insertion To confirm that the “chaperoning” effect of LDAO resulted from membrane destabilisation rather than solubilisation and stabilisation of the unfolded polypeptide chain by detergent monomers in the aqueous phase, we performed refolding trials analogous to those presented above in the presence of OG, which has a much higher CMC of >25 mM (Broecker and Keller 2013). In these experiments, we kept the OG concentration constant at 20 mM

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Fig. 3  Yield of OmpLA refolding into DLPC LUVs or SUVs destabilised by increasing concentrations of LDAO. OmpLA unfolded in 8 M urea was refolded by drop dilution at 50 °C to reach final concentrations of 0.33 mg/mL OmpLA, 0.87 M urea, 10.8 mM glycine, and 4 mM DLPC in the presence of different concentrations of LDAO, and the refolding mixture was agitated for 16 h at 50 °C. a SDS-PAGE of OmpLA refolded into DLPC LUVs in the presence of LDAO. As a reference, a sample without LDAO was boiled for

10 min at 97 °C (second lane). b Refolding yields in DLPC LUVs and SUVs as functions of LDAO concentration as obtained by densitometric quantification of gels such as that in a. Inset same data plotted against the LDAO/DLPC molar ratio in the bilayer, R, which was estimated by using the water-to-bilayer partition coefficient derived from the phase diagram (Fig. 2c). c Temperature dependence of refolding yield in the presence of 4 mM DLPC LUVs and 3.5 mM LDAO

and systematically increased the concentration of DLPC in the form of LUVs. As expected, the resulting decrease in the OG/DLPC molar ratio in the bilayer had a deleterious effect on refolding at 50 °C, which dropped from 62 % in the presence of only 2 mM DLPC to 39 % at a lipid concentration of 8 mM DLPC (data not shown). More interesting is the observation that high concentrations of monomeric OG in the aqueous phase were substantially less effective than much lower concentrations of LDAO (Fig. 3a, b) in catalysing membrane insertion, lending further credence to the implication of detergent-induced bilayer defects in enabling OmpLA to assume its native transmembrane topology. Besides membrane-bound detergent molecules, the protein itself might have an effect on bilayer integrity and, thus, permeability. Accordingly, it is conceivable that higher protein concentrations could facilitate refolding by creating more bilayer defects than would be caused by the detergent alone. To test this possibility, we compared the refolding kinetics at final OmpLA concentrations of 0.04 mg/mL (Fig. 4a) and 0.33 mg/mL (Fig. 4b) by quantifying the relative amount of refolded protein as a function of time. Refolding was essentially complete within 2 h at both concentrations but, on this relative scale, proceeded faster at the lower protein concentration tested (Fig. 4c).

Thus, even though the absolute amount of OmpLA inserted in a given time was larger at the higher concentration, it did not scale linearly with protein concentration, suggesting that formation of detergent-induced bilayer defects is the rate-limiting step in the process of coupled folding and insertion. Refolding into detergent‑destabilised long‑chain phospholipid bilayers Unlike relatively thin DLPC bilayers, thicker phospholipid membranes in the form of LUVs do not allow even partial refolding of OmpLA in the absence of chemical or molecular chaperones (Burgess et al. 2008). However, compromising the bilayer’s barrier function by controlled addition of subsolubilising amounts of LDAO afforded yields >95 % even on refolding at 50 °C into LUVs composed of the commonly used singly unsaturated long-chain phospholipid POPC (Fig. 5a). As was observed for DLPC, refolding improved monotonously with increasing detergent/lipid molar ratio in the bilayer; however, achieving essentially quantitative yields required substantially higher detergent contents approaching the maximum, saturating value (Fig. 5b). The latter amounts to RSAT = 1.39 at 25 °C (Textor et al. 2015) but, again, is expected to be somewhat

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Fig. 4  Kinetics of OmpLA refolding into LUVs. OmpLA unfolded in 8 M urea was refolded by drop dilution at 50 °C to reach final concentrations of a 0.04 mg/mL or b 0.33 mg/mL OmpLA, 0.87 M urea, 10.8 mM glycine, 4 mM DLPC, and 3.5 mM LDAO, and the refold-

ing mixture was agitated at 50 °C and monitored by SDS-PAGE. As a reference, a 300-min sample was boiled for 10 min at 97 °C. c Refolding yields as functions of time as obtained by densitometric quantification of the gels in a and b

Fig. 5  Refolding yield and enzymatic activity of OmpLA refolded from 8 M urea into POPC LUVs doped with increasing concentrations of LDAO. a SDS-PAGE of OmpLA refolded at 50 °C into 10 mM POPC LUVs in the presence of different concentrations of LDAO. As a reference, a sample without LDAO was boiled for 10 min at 97 °C. b Yield of OmpLA refolding into 10 mM POPC as a function of LDAO concentration as obtained by densitometric quantification of the gel depicted in a. Inset same data plotted against the LDAO/POPC molar ratio in the bilayer, R, which was estimated by assuming a water-to-bilayer partition coefficient of Kb/aq = 2.3 × 104 previously derived from a POPC/LDAO phase diagram at 25 °C (Tex-

tor et al. 2015). c Enzymatic activity of OmpLA refolded either into 4 mM POPC and 6 mM LDAO or into 10 mM POPC and 14.5 mM LDAO to a protein concentration of 0.33 mg/mL. Baseline-corrected absorbance at 412 nm, A, was plotted against time, t, after addition of CaCl2 at t  = 0. Final concentrations in the enzyme assay were, respectively, 32.4 or 3.92 µg/mL OmpLA, 393 or 119 µM POPC, 589 or 172 µM LDAO, 0.98 mM HEPC, 0.78 mM DTNB, and 19.6 mM CaCl2. Normalised specific activities of 12.7 and 14.4 µmol/(min mg) were derived from the average slopes of the kinetic traces at 32.4 and 3.92 µg/mL, respectively

higher at elevated temperatures. The proteoliposomes thus produced were enzymatically active (Fig. 5c); importantly, the absolute activity was proportional to the concentration of correctly folded OmpLA, suggesting that all of the refolded protein regained its native conformation and activity on LDAO-chaperoned insertion into POPC bilayer membranes. The normalised specific activity of OmpLA in LDAO-containing POPC membranes was ~14 µmol/

(min mg), approximately one-third lower than in DLPC bilayers but very similar to the value determined in the phosphocholine detergent DPC (Fig. 1b). Under subsolubilising conditions, phospholipid bilayers respond to monolayer curvature stress induced by the symmetric insertion of typical synthetic detergents such as LDAO, with clearly segregated headgroup and alkyl moieties, by acyl chain fluidisation and, consequently, lateral

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bilayer expansion and concomitant thinning (Karlovská et al. 2004; Nazari et al. 2012). The resulting decrease in lateral pressure in the acyl chain region and the reduction in membrane thickness are both expected to favour transient formation of aqueous bilayer defects, and the present findings demonstrate that this can be exploited to promote functional refolding of an integral membrane protein into preformed lipid bilayers. It should be noted that LDAO flip–flops rapidly between the bilayer leaflets of both DLPC and POPC membranes (data not shown), indicating that monolayer curvature strain resulting from symmetric detergent insertion is sufficient to induce the above effects. Nevertheless, it is conceivable that bilayer curvature strain caused by asymmetric accumulation of detergent in the outer vesicle leaflet could be equally or even more effective (Pantaler et al. 2000). Other classes of detergents may fulfil the same purpose, albeit through different mechanisms. In particular, detergents that do not mix well with phospholipids in the membrane are not distributed homogeneously within the bilayer but rather tend to cluster together in segregated domains (Nazari et al. 2012). Bilayer defects at the boundaries of such domains are believed to enhance membrane permeability (Heimburg 2007), similarly to the effect of coexisting gel and fluid domains near Tm (Danoff and Fleming 2015). Indeed, we have recently shown (Frotscher et al. 2015a, b) that a fluorinated octyl maltoside analogue termed F6OM, which partitions into POPC membranes but does not induce acyl chain disordering, also promotes the functional refolding and bilayer insertion of OmpLA. The domain-boundary mechanism seems to be less efficient in terms of refolding yield but more gentle in that the normalised specific activity of OmpLA refolded into POPC membranes with the aid of F6OM is about threefold higher than that observed here upon LDAO-mediated bilayer insertion; by contrast, the major advantage of our approach based on homogeneously disordering detergents such as LDAO is that it affords virtually quantitative refolding.

Conclusions In summary, we have shown that the controlled destabilisation of the barrier function of lipid bilayer membranes by judicious use of detergents can be exploited to promote the functional refolding and insertion of an integral membrane enzyme. On the basis of lipid/detergent phase diagrams, subsolubilising detergent concentrations can be chosen such as to increase membrane permeability by inducing monolayer curvature strain, acyl chain fluidisation, and membrane thinning. Without the need to solubilise the membrane, this enables the passage of hydrophilic protein moieties across the hydrocarbon core of the lipid bilayer,

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presumably by transient formation of aqueous bilayer defects. Acknowledgments  We thank Dr. Carolyn Vargas (University of Kaiserslautern) for helpful comments on the manuscript. This work was supported by the Deutsche Forschungsgemeinschaft (DFG) through International Research Training Group IRTG 1830 and by the Research Initiative BioComp. Conflict of interest  The authors declare no competing financial interest.

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