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
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Abstract |
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Keywords: BLM/modulation/mutants/patch clamp/porin
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Introduction |
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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, 2003). 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, 1978
). 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 currentvoltage relationships obtained when populations of channels are subjected to progressively ramped up and down voltages [see Baslé and Delcour (2004)
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., 1997; Phale et al., 1997
; Bainbridge et al., 1998b
). 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., 1996
; Phale et al., 2001
). 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, 1999
; Arbing et al., 2000
, 2001
). 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., 1995
; Müller and Engel, 1999
). 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, 1992). Acidic pH has been shown not only to affect the conductance and selectivity of OmpF (Benz et al., 1979
; Nestorovich et al., 2003
), but also to promote an increase in the frequency of closures (Buehler et al., 1991
; Todt et al., 1992
) 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., 2003
). 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., 2003
). 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, 1999
), 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.
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Materials and methods |
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Cells were grown in LuriaBertani 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, 1995; Guex and Peitsch, 1997
; Schwede et al., 2003
) using the wild-type structure as a template (access code 1OPF) (Cowan et al., 1995
). 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
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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Preparation of membrane fractions
Purification of outer membrane fractions was performed as described by sucrose density gradient ultracentrifugation (Delcour et al., 1989). 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., 2003), 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 SDSPAGE. 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., 1989). 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
. This pipette was filled with buffer A. Seals of 0.51.0 G
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). First, a hole of 50100 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) hexadecanehexane. 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 12 µg of pure porin into
4 ml of buffer in each compartment. For experiments where the insertion of a single trimer was desired, 1050-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., 2000). 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 2060 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).
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Results |
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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
L1,
L2,
L4,
L5 and
L8.
L6 and
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
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., 1996
), a region common to the wild-type and all the mutant proteins.
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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 currentvoltage 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 currentvoltage relationship reverses sign.
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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 L4 mutant in the positive voltage range only and between the wild-type and the
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
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., 1979; Buehler et al., 1991
; Todt et al., 1992
; Nestorovich et al., 2003
). Here we focused on the modulation of porin activity, as represented by the kinetics of openclosed 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., 2003
). 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., 2003
). The
L2,
L4,
L5 and
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
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
L1,
L7 and
L8 mutants were somewhat more dynamic at pH 7.0, but some variability in their behavior was encountered across bilayers (see
L1 and
L7 traces at pH 7.0, Figure 4B and D). The
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
L7 mutant (Figure 4D) and the
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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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., 2004
). Upward deflections from the BL at negative pipette voltages are transient closures and correspond to substates of the OmpF monomeric conductance (Baslé et al., 2004
). Only the
L4 mutant exhibits a somewhat enhanced gating to substates. However, even for this mutant, the open probability remains high.
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Discussion |
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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., 2001), 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)
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., 1994). 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., 1998a
; Robertson and Tieleman, 2002
).
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., 2003). We screened our mutants for this effect and found that the
L4,
L5 and
L6 mutants behave as wild-type, whereas the
L1,
L7 and
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)
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., 2003
). 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., 2004
). Conformational changes that lead to monomeric pore closure could very well be supported by other protein regions.
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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., 2003) have pointed to possible targets for further investigation.
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Acknowledgments |
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Received June 11, 2004; revised August 25, 2004; accepted September 28, 2004.
Edited by Hagan Bayley
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