Deletions of single extracellular loops affect pH sensitivity, but not voltage dependence, of the Escherichia coli porin OmpF

Arnaud Baslé, Randa Qutub, Mahsa Mehrazin, Jamie Wibbenmeyer and Anne H. Delcour1

Department of Biology and Biochemistry, University of Houston, 369 Science and Research Building 2, Houston, TX 77205-5001, USA

1 To whom correspondence should be addressed. E-mail: adelcour{at}uh.edu


    Abstract
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 Materials and methods
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 References
 
The molecular basis for the voltage and pH dependence of the Escherichia coli OmpF porin activity remains unknown. The L3 loop was previously shown not be involved in voltage dependence. Here we used seven OmpF mutants where single extracellular loops, except L3, were deleted one at a time. The proteins are expressed at levels comparable to wild-type and purified as trimers. Wild-type and mutant proteins were inserted into planar lipid bilayers for electrophysiological measurement of their activity. Current–voltage relationships show the typical porin channel closure at voltages greater than the critical voltage. Measurements of critical voltages for the seven deletion mutants showed no significant differences relative to wild-type, hence eliminating the role of single loops in voltage sensitivity. However, deletions of loops L1, L7 or L8 affected the tendency of channels to close at acidic pH. Wild-type channels close more readily at acidic pH and their open probability is decreased by ~60% at pH 4.0 relative to pH 7.0. For mutants lacking loop L1, L7 or L8, the channel open probability was found not to be significantly different at pH 4.0 than at pH 7.0. The other deletion mutants retained a pH sensitivity similar to the wild-type channel. Possible mechanistic scenarios for the voltage- and pH dependence of E.coli OmpF porin are discussed based on these results.

Keywords: BLM/modulation/mutants/patch clamp/porin


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The general diffusion porin OmpF of Escherichia coli has been the subject of intense investigation for several decades (Delcour, 2003Go; Nikaido, 2003Go). Owing to its large abundance in the outer membrane, the protein can be easily purified, which allowed for the determination of its crystal structure (Cowan et al., 1992Go) and the study of its pore properties by electrophysiology (Delcour, 2003Go). Structurally, the protein is characterized by the trimeric assembly of monomeric 16-stranded ß-barrels, each containing its own hydrophilic pore. Each barrel essentially spans the thickness of the membrane. Except for the third loop that folds inward and constricts the channel opening, seven long loops connect adjacent pairs of ß-strands on the extracellular side. The L2 loop projects somewhat sideways and is implicated in the thermal stability of the trimer (Phale et al., 1998Go). A residue at the tip of this so-called ‘latching loop’ (E71) forms stabilizing salt bridges with the R100 and R132 residues at the root of the L3 loop of the adjacent monomer (Phale et al., 1998Go). The constricting L3 loop participates in the size exclusion and selectivity properties of the pore. The roles of loops L1 and L4–L8 in channel function have remained undefined.

Protein channels are characterized by their ability to oscillate between two functionally distinct states: an ion-conducting open state and a non-ion-conducting closed state. General diffusion porins, such as OmpF, are able to fluctuate spontaneously between these states, although the open state is essentially predominant in conditions of neutral pH and low membrane potentials (Delcour, 2003Go). Although its functional significance is still unclear, the voltage dependence of porin activity has been established since the early studies of porin electrophysiology (Schindler and Rosenbusch, 1978Go). It is characterized by the step-wise closure of porin monomers above a threshold voltage, called the critical voltage (Vc). When populations of channels are investigated, the voltage-triggered closure can lead to an abrupt and drastic current reduction as many channels close at the same time. Interestingly, high voltages lead to stabilization of these closed states, since recovery from closures is not strictly reversible. This is seen in the form of typical hysteresis loops in the current–voltage relationships obtained when populations of channels are subjected to progressively ramped up and down voltages [see Baslé and Delcour (2004)Go for details].

The molecular basis for this drastic change in pore activity has been long sought. Elegant studies using tethering strategies to reduce or eliminate any possible motion of L3 across the pore have refuted the involvement of the L3 loop as a voltage effector (Eppens et al., 1997Go; Phale et al., 1997Go; Bainbridge et al., 1998bGoGo). The possibility that charged residues of L3 and the opposite barrel wall play the role of voltage sensors has been proposed on the basis that mutations in these residues render OmpF less voltage sensitive (Saint et al., 1996Go; Phale et al., 2001Go). Evidence for the involvement of extracellular loops in voltage dependence has been provided in the case of the major porin of Haemophilus influenzae type b, where chemical modification or site-directed mutagenesis of surface-accessible lysine residues of the L4 and L6 loops led to alterations in voltage sensitivity (Müller and Engel, 1999Go; Arbing et al., 2000Go, 2001Go). In the case of E.coli OmpF, there is some suggestion that extracellular loops might also be changing conformation upon voltage application. An atomic force microscopic study revealed that loops L1 and L4 to L8 form a somewhat compact structure that projects outwards about 13 Å away from the plane of the membrane and that this protrusion collapses into a 6 Å high doughnut-like structure when a high voltage is applied (Schabert et al., 1995Go; Müller and Engel, 1999Go). Taken together, all these studies on general diffusion porins of E.coli and H.influenzae hint at the possible role of extracellular loops in voltage sensitivity. This hypothesis was put to the test in the study presented here.

Another form of modulation with potential physiological relevance is the acidic pH-induced closure of OmpF. As microorganisms are likely to encounter environments of differing composition, the modulation of outer membrane permeability by pH might play an important role in adaptation and survival. Indeed, the inhibitory effect of acidic pH on porins is strong enough to lead to a decrease in overall outer membrane permeability (Todt and McGroarty, 1992Go). Acidic pH has been shown not only to affect the conductance and selectivity of OmpF (Benz et al., 1979Go; Nestorovich et al., 2003Go), but also to promote an increase in the frequency of closures (Buehler et al., 1991Go; Todt et al., 1992Go) and the appearance of closing events of size smaller than the full monomeric closures (also known as sub-conductance states or substates) (Nestorovich et al., 2003Go). Here also, the molecular basis for these effects of pH is not fully defined. Based on the fact that the D113C and E117C mutations produce conductance decreases of similar value to the differences observed between full monomeric state and the pH-induced substates, Bezrukov and colleagues intimated that the rapid substate fluctuations observed at acidic pH originate from the reversible protonation of the D113 and E117 residues of L3 (Nestorovich et al., 2003Go). They also present convincing evidence that these effects are electrostatic in nature, rather than structural. On the other hand, in the same study as mentioned for voltage dependence, Müller and Engel reported that the collapse of the extracellular protrusion is also seen at acidic pH (Müller and Engel, 1999Go), lending support that extracellular loops might alter their conformation to lead to pore closure when the pH is dropped.

In this study, we explored the possibility that deletions of single extracellular loops might affect the pH and/or voltage sensitivity of OmpF. We used a combination of site-directed mutagenesis, protein and membrane purification and electrophysiology to investigate the effect of these deletions on the kinetics of OmpF channels. The critical voltage for channel closure was found to be unchanged by these modifications. However, the deletion of L1, L7 or L8 rendered the channel less susceptible to closing with a drop in pH.


    Materials and methods
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Chemicals, media and buffer composition

Cells were grown in Luria–Bertani broth (1% tryptone, 1% NaCl and 0.5% yeast extract) with appropriate antibiotics (kanamycin at 100 µg/ml and tetracycline at 15 µg/ml) and 1 mM IPTG, as required. Tryptone and yeast extract were obtained from Difco Laboratories. n-Octyloligooxyethylene (octyl-POE) was purchased from Axxora. Other chemicals were obtained from Sigma or Fisher. For electrophysiology, the following buffers were used: buffer A (150 mM KCl, 10 µM CaCl2, 0.1 mM K-EDTA, 5 mM HEPES pH 7.2), buffer B (buffer A + 20 mM MgCl2), buffer G (1 M KCl, 5 mM HEPES, pH. 7.0), buffer H (buffer A with HEPES replaced by 5 mM MES, pH 5.6) and buffer I (1 M KCl, 5 mM MES, pH 4.0).

Mutant design and construction

Table I shows the amino acid sequences that were deleted in the construction of our mutants. The L3 loop has been studied extensively by others and shown not to be involved and was therefore omitted in our study. A deletion of 17 residues was originally performed for the L6 loop but the protein was not expressed (data not shown); therefore, we switched to the shorter 12-residue deletion presented here. A representation of the modeled structures is given for each deletion mutant in Figure 1. These structures were obtained by homology modeling of the mutated sequences by SWISS-MODEL via the ExPASy server of the Swiss Institute of Bioinformatics (Peitsch, 1995Go; Guex and Peitsch, 1997Go; Schwede et al., 2003Go) using the wild-type structure as a template (access code 1OPF) (Cowan et al., 1995Go). The deleted sequence of each loop is represented by a different color in the wild-type structure. The same color is used to highlight the four residues flanking the deleted sequence (two at the N-terminal side and two at the C-terminal side) that remain in each deletion mutant. The software was unable to model the last ß-strand in the mutant lacking L8. However, as shown below, the {Delta}L8 mutant is stable as a trimer and we therefore believe that the actual structure is that of a fully constructed ß-barrel.


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Table I. Deleted sequences

 


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Fig. 1. Molecular representations of the wild-type and deletion mutant OmpF structures. The wild-type structure is the published solved structure (Cowan et al., 1995Go). The structures of the deletion mutants were obtained by homology modeling, as described in Materials and methods. The deleted sequence of each loop is represented by a different color in the wild-type structure (L1, yellow; L2, blue; L4, orange; L5, green; L6, purple; L7, red; L8, cyan). The same color is used to highlight the four residues flanking the deleted sequence (two at the N-terminal side and two at the C-terminal side) that remain in each deletion mutant. The representations were made with the Visual Molecular Dynamics (VMD) program (Humphrey et al., 1996Go).

 
Mutations were introduced with a QuikChange mutagenesis kit (Stratagene) into the ompF gene cloned in the pNLF10 plasmid (Iyer et al., 2000Go). Mutations were identified by DNA sequencing. Subsequently the mutated gene was subcloned into a virgin plasmid using BamHI and SalI restriction enzymes and fully sequenced. Wild-type and mutant porins were expressed from the plasmid upon addition of 1 mM IPTG to AD100, an E.coli host that does not express ompF and ompC (Baslé et al., 2004Go). For protein purification, we used AD102, a strain that is missing not only ompF and ompC, but also ompA from the chromosome (Baslé et al., 2004Go). IPTG-dependent expression of ompF was verified by colicin-A sensitivity and western blotting. The ompF gene with an L2 loop deletion was obtained from R.Koebnik (Phale et al., 1998Go) and subcloned into pNLF10.

Preparation of membrane fractions

Purification of outer membrane fractions was performed as described by sucrose density gradient ultracentrifugation (Delcour et al., 1989Go). Protein concentration was determined with the bicinchoninic assay (Pierce).

Protein purification

Purification of wild-type and mutant OmpF protein was essentially done as described for Vibrio cholerae porins (Simonet et al., 2003Go), except that protein extraction with the detergent octyl-POE was done at 37°C instead of 4°C and three extractions at 3% were performed instead of two. Purification of wild-type and mutant OmpF protein was performed by anion-exchange chromatography (Mono Q HR10/10, Pharmacia) and the protein eluted between 250 and 400 mM NaCl (in 0.5% octyl-POE, 10 mM Na phosphate buffer, pH 7.6). Subsequently, OmpF-containing fractions were further purified by size-exclusion chromatography on a Hiload 26/20 Superdex 200 Prep grade column (Pharmacia) in 0.5% octyl-POE, 50 mM NaCl, 10 mM Na phosphate buffer, pH 7.6. Proteins were identified by western blotting. Protein visualization and purity were assessed by silver staining after SDS–PAGE. Samples were either left at room temperature or heated at 96°C for 10 min prior to electrophoresis. Pure OmpF was kept at –80°C in 0.5% octyl-POE, 10 mM Na phosphate buffer, pH 7.6 and 50 mM NaCl, prior to use in electrophysiology. Protein concentration was determined with the bicinchoninic assay (Pierce).

Reconstitution into liposomes and patch clamp electrophysiology

Reconstitution of outer membrane fraction was performed into soybean phospholipids as described (Delcour et al., 1989Go). The large multilamellar liposomes thus formed were placed into buffer B in the patch clamp chamber (see buffer composition above). The presence of magnesium causes the collapse of the liposomes followed by the emergence of large unilamellar blisters that can easily be sampled by patch clamp. A borosilicate glass capillary (Drummond No. 2-000-100) was pulled into a pipette whose tip diameter gave a resistance of ~10 M{Omega}. This pipette was filled with buffer A. Seals of 0.5–1.0 G{Omega} were formed by bringing the pipette tip into contact with the blister membrane. After seal formation, the patch was momentarily exposed to air and brought back to the buffer (excised patch). This ensured that there was only a single bilayer of membrane at the tip of the pipette. The bath was then perfused with buffer A or H.

Reconstitution into bilayers and planar lipid bilayer electrophysiology

Virtually solvent-free planar lipid bilayers were formed by the technique outlined by Montal and Mueller (1972)Go. First, a hole of 50–100 µm is made in a 10 or 25 µm thick Teflon film (Goodfellow) using a high-frequency spark tester generating an electrical arc (Type PPM MK3, Buckleys). The size and the shape of the hole were checked using a dissecting microscope. The film was then sandwiched between two half glass cells and the hole was treated with a mixture of 1:40 (v/v) hexadecane–hexane. After 45 min, the chamber was filled with buffer on both sides and 30 µl of lipid were deposited on the surface of the buffer. Lipid was prepared from azolectin II-S from soybean (Sigma) dissolved in hexane at a concentration of 0.5% (w/v). Bilayers were formed about 10 min after lipid addition, by lowering the buffer below the hole and then raising the buffer again above the hole. Reconstitution of channels was performed by adding 1–2 µg of pure porin into ~4 ml of buffer in each compartment. For experiments where the insertion of a single trimer was desired, 10–50-fold less protein was used. The choice of buffer was dictated by the experimental design. Voltage ramps were applied with an Agilent function generator (Model 33120A) at a rate of 1.6 mV/s. The critical voltage for voltage-dependence (Vc) was defined as the highest voltage at which the slope of the curve tangent reverses sign.

Data acquisition and analysis

The voltage across the membrane was clamped at different values using an Axopatch 1D amplifier (Axon Instrument). For bilayer experiments, the cis side of the membrane was defined as ground, as documented by others (Van Gelder et al., 2000Go). The CV-4 headstage and the CV-4B headstage were used for patch clamp experiments and for bilayer experiments, respectively. The resulting current was filtered at 1 kHz, digitized at 84.75 or 100 µs sampling intervals for 20–60 s traces (ITC-18, Instrutech). Longer recordings (up to 10 min) were digitized at 1.25 ms sampling intervals. Traces acquired on a PC (Acquire, Bruxton) were also recorded on charts (Graphtec). Finally, the data were analyzed using a program specifically developed in the laboratory using Microsoft Visual Studio C.net. The open probability Po was calculated as the ratio of the observed integrated current obtained over a 10 min long recording to the total current expected for the same duration if the current value remained at the fully open level (macroscopic current).


    Results
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 Abstract
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 Materials and methods
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Protein expression and purification

In order to investigate the role of OmpF extracellular loops in voltage-gating and pH dependence, we compared the electrophysiological behavior of wild-type OmpF with that of the seven deletion mutants described in Materials and methods. For this, we used both patch clamp electrophysiology on liposomes containing reconstituted membrane fractions and planar lipid bilayers containing reconstituted purified proteins. Figure 2A shows that all mutant proteins are expressed at levels comparable to wild-type levels. The bands at ~39 kDa represent the OmpF monomer and the relative positions of the mutant bands are visibly lower for {Delta}L1, {Delta}L2, {Delta}L4, {Delta}L5 and {Delta}L8. {Delta}L6 and {Delta}L7 migrate at about the same height as wild-type OmpF. The band at ~30 kDa represents OmpA, which for an unknown reason was not expressed at a detectable level in the strain AD100/pNLF10{Delta}L5. The band assignments were confirmed by the western blot shown in Figure 2B, which was obtained with the F4 antibody specifically directed to the L3 loop of OmpF (Simonet et al., 1996Go), a region common to the wild-type and all the mutant proteins.



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Fig. 2. Expression level and purity of the wild-type and loop deletion mutants of OmpF. (A) Coomassie Brilliant Blue-stained SDS–PAGE gel of outer membrane fractions obtained from strains expressing the wild-type (wt) and mutant channels as indicated. All samples were boiled at 96°C for 10 min prior to loading on a 12% gel. The gel was stained with Coomassie Brillant Blue G-250. The bands corresponding to OmpF and OmpA are labeled F and A, respectively. (B) Western blot of a similar gel as above using the F4 antibody for immunodetection. The band corresponding to OmpF is labeled F. (C) Silver-stained SDS–PAGE gel of purified proteins obtained from strains expressing the wt and mutant channels as indicated. Samples were either left at room temperature (lanes marked with a – sign) or were boiled for 10 min at 96°C (lanes marked with a + sign) prior to loading on a 12% gel. The leftmost lane contains the molecular weight markers. The bands corresponding to the trimeric and monomeric forms are labeled T and M, respectively. For panels (A) and (B), only the relevant part of the gel is shown (30–45 kDa); for panel (C), the entire gel is shown to document sample purity.

 
We also purified wild-type and mutant OmpF, as described in Materials and methods. Figure 2C shows a silver-stained SDS–PAGE gel, where each pure protein has been loaded as such or after a 10 min incubation at 96°C to denature the trimers. Non-heated samples run as a ladder of high molecular weight bands, as typical for OmpF trimers with various amounts of associated lipopolysaccharides (Fourel et al., 1994Go). In addition, some dimer form is evident for the {Delta}L1 and the {Delta}L2 mutants, probably because of the increased lability and possibly because of some SDS sensitivity of these mutants (Phale et al., 1998Go). {Delta}L1-carrying strains also exhibit a slower growth and the {Delta}L1 mutant is somewhat harder to study by electrophysiology. Finally, the {Delta}L4 sample showed some proteolytic degradation, but only when the sample was heated since the <30 kDa bands are absent from the non-heated sample.

Voltage-induced inactivation

General-diffusion porins, such as OmpF, OmpC and PhoE, are characterized by a tendency to close upon high transmembrane potentials. Because this type of closure is distinct from the activity seen when channels spontaneously oscillate between open and closed states, we refer to this phenomenon as voltage-induced inactivation. A hallmark of this behavior is that it requires a threshold voltage, called the critical voltage (Vc), and displays a typical hysteresis pattern when voltages are ramped up and down across a membrane bilayer.

Figure 3A shows a typical current–voltage plot, obtained from wild-type OmpF. The data were obtained when the system equilibrated after purified proteins, added to both sides of the membrane, were incorporated into a planar lipid bilayer. It is estimated that about 170 trimers have inserted. Then, the bilayer was subjected to consecutive voltage ramps of 1.6 mV/s from 0 to 200 mV, then down to –200 mV, to finally back to 0 mV. As the voltage increases from 0 to 200 mV, the current increases as expected owing to the progressively larger driving force for ion movement (ohmic behavior). When the voltage reaches ~130 mV, the current decreases and continues to do so despite the larger and larger potentials. This non-ohmic behavior is indicative of a decrease in the number of open channels, i.e. channel closure. This pattern is reproduced in the negative voltage range. The critical voltage Vc is determined as the voltage at which the slope of the current–voltage relationship reverses sign.



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Fig. 3. Voltage dependence of wild-type OmpF and the loop deletion mutants. (A) A typical macroscopic current–voltage (IV) curve was obtained for OmpF in BLM in buffer G (1 M KCl, 5 mM HEPES, pH. 7.0), by ramping the voltage between +200 and –200 mV at 1.6 mV/s (sequence shown by the arrows near the trace). (B) The critical voltages (Vc) were obtained from IV plots as the highest voltage at which the tangent to the curve reverses sign (open bars, positive voltage regime; hatched bars, negative voltage regime). The graph shows the averages and s.e.m. obtained from 8–19 experiments.

 
Voltage ramps are widely used in the porin literature to document voltage dependence. Although the ramp rate might affect the value of Vc, we are not aware of any systematic analysis of this parameter in the literature. We chose a rate of 1.6 mV/s in accordance with other groups (Lakey and Pattus, 1989Go; Bredin et al., 2002Go), as this is an experimental condition that gives a reproducible quantitative measurement of Vc (as seen from the small error bars in Figure 3B). Since the voltage dependence of wild-type and mutant channels is studied with the same voltage ramp rate, the results obtained with the different proteins are directly comparable.

Figure 3B shows the averages and standard errors of the mean (s.e.m.) of Vc obtained in the positive and negative voltage ranges for wild-type OmpF and the seven deletion mutants. The average Vcs were significantly different (P < 0.05, t-test) between wild-type and the {Delta}L4 mutant in the positive voltage range only and between the wild-type and the {Delta}L7 mutant in the negative voltage range only. These differences were small (<15%) and not reproduced in both voltage regimes. Significant differences were seen between the wild-type channel and the {Delta}L6 mutant in both the positive and negative voltage ranges. These differences were also rather mild (~15% increase). Therefore, we conclude that the deletion of single extracellular loops does not appreciably affect the voltage-dependence of OmpF.

Sensitivity to acidic pH

The sensitivity of various features of OmpF channels to acidic pH has been documented by many (Benz et al., 1979Go; Buehler et al., 1991Go; Todt et al., 1992Go; Nestorovich et al., 2003Go). Here we focused on the modulation of porin activity, as represented by the kinetics of open–closed transitions and the probability that the channel is in its open state (open probability Po). In a very thorough study, Nestorovich and co-workers studied the dependence of OmpF activity over a wide pH range (Nestorovich et al., 2003Go). Their data helped us choose pH 4.0 as the pH for comparison with pH 7.0 in our investigation of wild-type and mutant OmpF channels. This acidic pH was chosen because it was the highest pH that gave a clear modulatory effect without being too harsh for the bilayer and the proteins. For this study no more than two trimers were inserted in the bilayer at a time.

Figure 4 shows traces of wild-type and some mutants that were recorded in BLM at the indicated voltages for 10 min. The vertical line at the beginning of each trace represents either a trimer insertion or the current deflection due to the voltage step from 0 to 100 mV. At neutral pH and at 100 mV (below Vc), the wild-type channel remains essentially fully open for the whole 10 min recording (Figure 4A). Under these conditions, there is very little tendency for the channel to close since the voltage is below the critical voltage. Hence the open probability of the wild-type channel is close to 1 at pH 7.0 (see below). At pH 4.0, the wild-type channels close more readily and they spend slightly less than half of the 10 min recording in the open state (thus the open probability drops to ~40%; see below). This type of behavior is similar to that reported by Bezrukov and colleagues, who document a faster and faster onset of closure at more and more acidic pHs (Nestorovich et al., 2003Go). The {Delta}L2, {Delta}L4, {Delta}L5 and {Delta}L6 mutant channels behaved as the wild-type and were affected by acidic pH. These proteins were easily closed at pH 4.0, as shown for the {Delta}L4 mutant in Figure 4 (other mutants are not shown). In other words, the deletion of the L2, L4, L5 or L6 loop does not seem to affect the pH sensitivity of the channel. The {Delta}L1, {Delta}L7 and {Delta}L8 mutants were somewhat more dynamic at pH 7.0, but some variability in their behavior was encountered across bilayers (see {Delta}L1 and {Delta}L7 traces at pH 7.0, Figure 4B and D). The {Delta}L1 mutant (Figure 4B) showed a great deal of fast transient closures at pH 4.0, but was less prone to permanent closure at pH 4.0 than at pH 7.0. The {Delta}L7 mutant (Figure 4D) and the {Delta}L8 mutant (not shown) behaved in a similar way at pH 7.0 and 4.0 and remained fairly open during the 10 min recording at both pHs.



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Fig. 4. pH sensitivity of wild-type and OmpF mutants in BLM experiments. The 10 min current traces were obtained at 100 mV at the indicated pH in buffer G (1 M KCl, 5 mM HEPES, pH. 7.0) or I (1 M KCl, 5 mM MES, pH 4.0), from purified channels incorporated into BLM. The dashed lines show the zero current level for each trace. The vertical line represents the change in current obtained either when an insertion occurs or when the voltage was switched from 0 to 100 mV. Downward deflections from the trace represent closures. The sample interval is 1.25 ms.

 
The quantitative analysis of the pH effect on OmpF and the loop deletion mutants is presented in Figure 5. The graph shows the average and s.e.m. for the open probability of each porin at pH 7.0 and 4.0, calculated for 10 min long recordings such as those in Figure 4. In some cases, the values of pH 7.0 and 4.0 originate from the same bilayer that sustained the perfusion of buffer. However, in most cases, the values were obtained from independent experiments. A statistical analysis shows that significant differences in open probability were found between pH 7.0 and 4.0 conditions for wild-type, the {Delta}L2, {Delta}L5 and {Delta}L6 mutants (P < 0.05, t-test), but not for the {Delta}L1, {Delta}L7 and {Delta}L8 mutants. A significant difference in open probability was found for the {Delta}L4 mutant at the P < 0.1 level. Hence we conclude that the deletion of single loops L1, L7 and L8 diminishes the pH sensitivity of OmpF.



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Fig. 5. Effect of acidic pH on the open probability of the wild-type and mutant OmpF porins. The open probability was calculated from 10 min long recording of porin activity in BLM at pH 7.0 (open bar) (buffer G: 1 M KCl, 5 mM HEPES, pH 7.0) and at pH 4.0 (hatched bar) (buffer I: 1 M KCl, 5 mM MES, pH 4.0) at 100 mV. The average values were obtained from the number of experiments shown at the top of the bars. The error bars are s.e.m. values.

 
As reported by Bezrukov and colleagues (Nestorovich et al., 2003Go), we also found an increase in the amount of noise in the traces when the channels are investigated at pH 4.0. This noise is due to the rapid transitions of channels to states that are not fully closed (called substates or sub-conductance states). Transitions to these states occur at pH 7.0 (Baslé et al., 2004Go), but are increased in frequency in acidic pH. This was particularly true for the {Delta}L1 mutant. These events are extremely transient and, despite their frequent occurrence, they do not greatly affect the overall open probability, whose value is mostly dictated by the slow kinetics of full monomeric closures, such as those seen on the traces obtained at pH 4 for the wild-type and {Delta}L4 mutant, for example (Figure 4).

Patch-clamp experiments were used to obtain a more detailed view of the electrophysiological signature of the mutants, in particular with respect to these substates. No significant differences in the kinetics of spontaneous transitions were found between the wild-type and the mutant channels at pH 7.0 at –60 mV. Hence representative patch clamp traces are shown only for wild-type and some mutants (Figure 6). The macroscopic current is denoted ‘baseline’ (BL) and represents the current flowing through all the open pores of the patch. For theses traces, the macroscopic current was ~–70 pA at –60 mV, indicating that only one trimer was present in each patch (Baslé et al., 2004Go). Upward deflections from the BL at negative pipette voltages are transient closures and correspond to substates of the OmpF monomeric conductance (Baslé et al., 2004Go). Only the {Delta}L4 mutant exhibits a somewhat enhanced gating to substates. However, even for this mutant, the open probability remains high.



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Fig. 6. pH sensitivity of wild-type and some OmpF mutants in patch clamp experiments. The 20 s long traces were obtained for the indicated proteins from reconstituted outer membrane fractions at pipette voltages of –60 mV. For each protein, the traces originate from the same patch sequentially exposed to buffer A (150 mM KCl, 10 µM CaCl2, 0.1 mM K-EDTA, 5 mM HEPES, pH 7.2) and buffer H (150 mM KCl, 10 µM CaCl2, 0.1 mM K-EDTA, 5 mM MES, pH 5.6). The preferred current level, denoted baseline (BL), corresponds to the total current flowing through all open porins. Upward deflections from baseline represent transient closures.

 
The effect of pH was studied in patch clamp by perfusing the bath with a solution identical with the patch clamp buffer typically used, but at pH 5.6 (buffer H, see Materials and methods). These experiments were performed before the publication by Bezrukov's group and the pH was originally chosen for consistency with prior patch clamp experiments on OmpC (Liu and Delcour, 1998Go). In addition, the patch clamp traces were recorded for only 20 s and therefore mostly document the changes in the fast kinetics of substate gating. The trends shown by these traces, however, are consistent with the stronger effect observed in bilayer experiments at pH 4.0. A moderate increase in substate gating and monomeric closures is observed for wild-type OmpF (Figure 6), in accordance with the mild effect documented by others (Nestorovich et al., 2003Go). We have found considerable variability in the pH response of the {Delta}L4 mutant. About 50% of the {Delta}L4 patches appear unaffected by pH 5.6 (Figure 6C), whereas the other half showed a pronounced increase in spontaneous closures (Figure 6B). The reasons for these apparent two populations are unknown. However, this trend is also evident in bilayer experiments and led to the somewhat larger error bars for this mutant in Figure 5. In agreement with the bilayer results, the {Delta}L7 mutant appears to have lost sensitivity to pH. This was also the case for the {Delta}L8 mutant (data not shown).


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 References
 
OmpF is a ß-barrel pore-forming protein containing eight extracellular loops. One of these, L3, is folded into the barrel and acts as a constriction loop inside the pore. Previous studies have failed to demonstrate a major role for L3 in voltage-induced pore closure (Saint et al., 1996Go; Eppens et al., 1997Go; Phale et al., 1997Go). Perhaps this is not surprising since a toxin such as {alpha}-hemolysin, also a ß-barrel, lacks a constriction loop, but is still capable of voltage gating (Korchev et al., 1995Go; Song et al., 1996Go). The idea that the constriction loop might be a voltage-dependent gate was appealing, but must be put aside.

The next most attractive hypothesis is to involve the extracellular loops in voltage-dependent conformational changes leading to decreased ion flow. Our approach has been to delete each loop one at a time, but these types of manipulations also left the mutant OmpF with normal voltage dependence. Based on these results, it is tempting to rule out extracellular loops as voltage effectors as well. Indeed, since differences in voltage sensitivities can be found upon single amino acid changes in the loops of H.influenzae porin (Arbing et al., 2001Go), it would be expected that a loop deletion should have an effect, if this loop indeed participates in voltage gating, even if it is in concert with other loops. This might very well be the case, but caution has to be exercised, as there might be redundancy in loop function. The AFM work of Müller and Engel (1999)Go documented a conformational change of a fairly large extracellular domain of OmpF and it would therefore be expected that many loops might be involved. It is possible that the deletion of a single one is not sufficient to impair the movement of the others or to bring about detectable changes in critical voltages.

If extracellular loops and L3 are not involved, alternative hypotheses for molecular mechanisms of closure are changes in the ß-barrel structure or in the intrinsic electrostatic field shown to exist at the constriction of the pore (Karshikoff et al., 1994Go). The ß-barrel is unlikely to undergo large structural changes that could lead to pore closure due to the large number of hydrogen bonds between adjacent strands. Perhaps the applied field perturbs the intrinsic potential in such a way that electrostatic effects on permeating ions hamper their movement through the pore, which would resemble a closing event in electrophysiological traces. These ideas have been proposed by others (Bainbridge et al., 1998aGo; Robertson and Tieleman, 2002Go).

In contrast to voltage dependence, our results suggest that some extracellular loops may play a role in pH sensitivity. Wild-type OmpF was reported to close more readily at 100 mV when the pH is 3.9 than when it is neutral (Nestorovich et al., 2003Go). We screened our mutants for this effect and found that the {Delta}L4, {Delta}L5 and {Delta}L6 mutants behave as wild-type, whereas the {Delta}L1, {Delta}L7 and {Delta}L8 mutants lost their pH sensitivity. It is striking that the L7 and L8 loops each carry four acidic amino acids that can be protonated at acidic pH, a number much larger than for any other extracellular loop. The loss of pH sensitivity in these loop deletion mutants may be due to the removal of these residues. As shown in Figure 7, the L1, L7 and L8 loops are also in close proximity to each other, forming a distinct domain from the other loops. The AFM study of Müller and Engel (1999)Go suggested that the extracellular loops of OmpF undergo a reversible conformational change at pH. 3.0. Whether the loops are involved in pH sensing or in participating in the pH-induced conformational changes (pH gating) cannot be distinguished from our results. In their study of OmpF pH dependence, Nesterovitch and colleagues argue that the observed pH effects on conductance, selectivity and substate gating may be due to the titration of pore residues D113 and E117 (Nestorovich et al., 2003Go). Our results do not necessarily refute this argument, as there may be multiple sites for pH sensitivity. Our study has focused specifically on changes in open probability, which is mostly determined by the slow kinetics of monomeric closures. Although visual inspection of the traces presented in Nesterovitch et al.'s study clearly shows that open probability is affected by pH, this parameter was not included in the titration curves that support the role of D113 and E117 presented by these authors. The involvement of these residues of the constriction zone makes a lot of sense when one considers the features of porin activity that are typically associated with pore properties, i.e. conductance, selectivity and even substate gating, which we believe involves the L3 loop (Baslé et al., 2004Go). Conformational changes that lead to monomeric pore closure could very well be supported by other protein regions.



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Fig. 7. Location of acidic residues on the extracellular loops of an OmpF monomer. (A) View from the extracellular side; (B) side view. Acidic residues of extracellular loops are shown as CPK models in magenta for loops L2, L4, L5 and L6 and in turquoise for loops L1, L7 and L8. The structures were represented using RasMol (Sayle and Milner-White, 1995Go).

 
It is interesting that the {Delta}L2 mutant was more sensitive than wild-type to pH, with a 75% reduction in open probability at pH 4.0 compared with the 58% reduction for wild-type OmpF. Perhaps this might relate to the fact that the L2 residues E71 and D74 might contribute to the electrostatics of the channel, as suggested by the work Karshikoff's and Jakobsson's groups (Karshikoff et al., 1994Go; Varma and Jakobsson, 2004Go), since the L2 loop projects sideways and dips into the pore of the adjacent monomer. Alternatively, this enhanced sensitivity might relate to the enhanced instability of the trimer that is introduced by deletion of the latching loop. Whatever conformational changes are triggered by acidic pH to bring about pore closure might be favored in a more plastic channel that has lost some of its associations with its neighbors.

In conclusion, our study has provided some clues on the molecular determinants that underlie two major forms of modulation of OmpF porin activity by transmembrane voltage and acidic pH. The nature of the events that lead to voltage-induced porin closure remains ill-defined and will continue to pose a challenge. Most likely, biophysical approaches that combine electrophysiology with accessibility studies to modifying agents or spectroscopic analyses of conformational changes will need to be used. This may also be the case for a deeper understanding of the pH-induced changes, although our work and that of others (Nestorovich et al., 2003Go) have pointed to possible targets for further investigation.


    Acknowledgments
 
We are grateful to Ralf Koebnick for providing the plasmid with the {Delta}L2 ompF mutant and Jean-Marie Pagès for the gift of the F4 antibody. Valérie Simonet is acknowledged for her help in the cloning procedures. The work was supported by NIH grant AI34905.


    References
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
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Received June 11, 2004; revised August 25, 2004; accepted September 28, 2004.

Edited by Hagan Bayley





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