I. INTRODUCTION

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Cells experience a wide variety of mechanical stimuli ranging from thermal molecular agitation to potentially destructive osmotic pressure gradients. Therefore, from the onset, living organisms faced a basic dilemma in their evolution. As a first priority, they required mechanisms that would protect their delicate cell membrane from potentially damaging mechanical stimuli. However, to interact with their changing mechanical environment (e.g., during feeding, escaping, or mating), they needed mechanosensitivity. Different organisms have solved the dilemma by different strategies. Bacteria and plants evolved a rigid cell wall that protects their plasma membrane from excessive dilation. However, with this strategy they sacrificed not only cell deformability but also mechanosensitivity. In contrast, animals have adopted strategies that protect their cell membrane while preserving a high degree of cell deformability and mechanosensitivity.

The first mechanosensitive (MS) processes may have evolved as backup mechanisms for cell protection. For example, a large nonselective membrane pore activated by osmotic swelling will release the cell's contents and thereby reduce intracellular pressure and membrane tension. Similarly, tension-sensitive fusion of intracellular membrane vesicles with the cell membrane will act to reduce bilayer tension. These basic mechanisms of mechanically gated (MG) channels and MS exocytosis may have been subsequently refined to participate in cell signaling. For example, MG channels and/or MS transmitter release are implicated in a myriad of physiological processes, including touch and pain sensation (46, 292, 403), hearing and vestibular function (148, 190), blood pressure control (45, 61), salt and fluid balance (32), micturition (36), tissue growth (98), cell volume regulation (301, 308, 430), and turgor control (147, 265). Furthermore, abnormalities in these mechanisms may contribute to neuronal (93) and muscular degeneration (116), cardiac arrhythmia (86, 117, 162), hypertension (224), arteriosclerosis (90), and glaucoma (282).

The external mechanical forces that dominate a cell vary depending on its size and relationship with other cells. For example, unicellular organisms like Escherichia coli are constantly jostled by the forces of Brownian motion that tend to keep them in suspension. In contrast, multicellular organisms require specific MS mechanisms that constantly adjust their position in response to gravity. Furthermore, specific cells, depending on their location within an organism and association with ancillary structures, may be selectively exposed to specific forms of mechanical stimuli, including steady indentations, high-frequency vibrations, osmotic pressure gradients, and hemodynamic pressure and fluid shear stresses. All external stimuli act on top of a dynamic background of various internally generated forces (e.g., arising from hydrostatic pressure, cytoskeletal polymerization, and molecular motors) that are important in determining cell shape, growth, mobility, and adhesion (15, 201). To monitor and respond selectively to these different forces most likely requires multiple, parallel signaling pathways, with each pathway designed to extract specific information regarding the "relevant stimulus" while filtering out irrelevant stimuli.

Over the last 20 years, the molecular nature of specific MS membrane processes has been identified. These include MG membrane ion channels (156, 265, 286, 350) and MS receptors (53, 320), enzymes (241, 270), intracellular Ca²⁺ release (204) and transmitter release (63). Because each of these elements or processes may interact with one another, as well as with other non-MS elements, difficulties can arise in distinguishing mechanisms that are directly or indirectly affected by mechanical forces. Furthermore, given that a single cell may express multiple mechanotransducers, a challenge can arise in determining which transducer mediates a specific MS function (i.e, cause and effect relations). A notable example is vertebrate tactile sensation where the basic distinction between physical and chemical mechanisms of mechanotransduction has yet to be made (cf. Refs. 135, 292). In principle, one should be able to identify the mechanotransducer by comparing its specific properties (i.e., sensitivity, kinetics, and pharmacology) with those of the MS function.

The membrane of most animal cells is a composite structure of extracellular (EC), bilayer, and cytoskeletal (CSK) layers. Because of its integrated nature, any externally applied force produces varying tensions and strains in multiple elements within the three layers (200). For this reason, it becomes problematic in identifying a single membrane property that may be directly involved in the mechanotransduction process. To overcome this problem, we adopt a hierarchical approach and consider a variety of membrane preparations, progressing from the simple artificial bilayer vesicle to increasingly more complex cells (i.e., from bacteria to animal cells). The rationale for this approach is that if characteristics of a particular mechanism can be identified (i.e., "finger-printed") in a simple system, one should be better positioned to recognize its operation in more complex systems. Our approach would seem justified by the reoccurring theme in evolution in which basic mechanisms that first evolved in prokaryotes are conserved and refined to carry out more diverse and specialized functions in eukaryotes. However, some processes such as exocytosis/endocytosis and release of Ca²⁺ from intracellular membrane stores are unique to eukaryotes (54), and therefore, their mechanosensitivity must reflect more recently acquired mechanisms of mechanotransduction.

	II. LIPID BILAYER STRUCTURE AND ITS RESPONSE TO MECHANICAL DEFORMATION

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Because the bilayer is the core structure around which all other membrane components are arranged, it is critical to understand its molecular packing and how this packing may change under steady state and dynamic mechanical deformation. For example, intrinsic delays or relaxations in the response of the bilayer to deformation may be reflected in the functional dynamics (i.e., frequency response and adaptive behavior) of MG channel activities. The bilayer is composed of lipid molecules that form two monolayers stabilized by van der Waals forces and the "hydrophobic" effect between the "hidden" acyl lipid chains. In addition, water molecules surrounding each lipid headgroup form hydrogen bonds that further stabilize the bilayer (180). Water molecules also penetrate deeper into the bilayer, hopping between acyl chain packing defects, such as trans-gauche kinks. For example, it is estimated that ~4,000 water molecules pass a single phospholipid per second compared with 1 Na⁺ every 70 h (85). At reduced temperatures, lipid bilayers undergo a liquid-gel phase transition in which the acyl chain packing becomes more ordered (187a). Furthermore, bilayers made up of more than one phospholipid can undergo lateral phase separations (138, 221). In the case of cell membranes, it is generally assumed that their complex lipid and cholesterol makeup ensure a highly fluid state at physiological temperatures. However, recent studies indicate preferential packing of sphingolipids and cholesterol into floating platforms or rafts of lipid that can be isolated as detergent-insoluble membrane complexes (369a). Although the occurrence of such phase separations will complicate bilayer and cell membrane mechanics, their specific effects have been little studied.

In principle, it is possible to study the mechanics of planar lipid bilayers, typically formed by painting a film of phospholipid over a hole in a plastic barrier, (e.g., see Ref. 112). However, such studies are complicated by the presence of a torus of disordered lipid that can act as a lipid reservoir for formation of new bilayer (336, 433a). In contrast, the lipid vesicle is a more "cell-like" structure in that it is a closed system that has its volume set by the osmotic activity of the aqueous environment. How the lipid vesicle responds to mechanical deformation depends on both extrinsic (e.g., size and geometry) as well as intrinsic properties (i.e., material elastic properties) (see Refs. 107, 109). For example, a deflated vesicle filled with volumes insufficient to form a sphere is highly deformable but somewhat unstable with a tendency to spontaneously "bud" or vesiculate. In contrast, a vesicle inflated by osmotic or hydrostatic forces into a sphere is stable but shows limited deformability in that with further inflation (i.e., 2-4%) it either ruptures or under specific circumstances forms pores (452). Transient pore formation by releasing intravesicular pressure will preserve vesicle structure and thus may have served as an inbuilt protection mechanism for primordial cells before they evolved protein mechanisms.

With the assumption that the lipid bilayer behaves as an elastic solid, its intrinsic mechanical properties can be characterized by four elasticity constants (or moduli) that describe the response of a unit area of bilayer to compression, expansion, bending, and extension (108, 109). The larger the moduli, the greater the resistance to that form of deformation. Elastic deformations are directly proportional to and follow instantaneously the application and removal of external forces. In comparison, viscoelastic or plastic deformations show time dependence, and one has to take into account the different viscosity coefficients for each type of deformation (108). The elastic moduli of bilayer vesicles and human red blood cells (RBCs) have typically been measured with the micropipette aspiration technique (106). A critical feature of this technique is that there is minimal membrane adherence to the pipette to ensure reversible and unimpeded movement of the aspirated portion of the membrane within the pipette. Under these circumstances, the membrane tensions developed during aspiration can be assumed isotropic throughout the vesicle, with the membrane protrusion drawn into the pipette serving as an amplified measurement of membrane area changes. In contrast, in patch-clamp recording, the membrane adheres tightly to the walls of the pipette (153). As long as the membrane-glass adhesion is not disrupted (i.e., the patch boundary remains constant), tension changes will be restricted to the "free" membrane area that spans the inside of the pipette (see sect. VIIIA).

An early study based on the effects of pressure on the lipid bilayer phase transition demonstrated that hydrostatic pressures up to ~1 × 10⁷ N/m² (i.e., 100 atmospheres) did not significantly alter the bilayer density change associated with the phase transition (378). Based on this result, Evans and Hochmuth (108) estimated the bilayer compressibility modulus was between 10⁹ and 10¹⁰ N/m², similar to that of most "incompressible" fluids. A subsquent study based on X-ray diffraction analysis of osmotically stressed liposomes gave similar estimates (275, see also Ref. 322). Although higher compressive forces (i.e., 500-1,500 atm) have since been shown to increase acyl chain packing density and squeeze water out of pure phospholipid bilayer, these effects are minimized in bilayers that include cholesterol (16, 334a). Thus one may assume that the bilayer of the cell membrane is volumetrically incompressible and will maintain a constant density during the mechanical deformations encountered under physiological conditions. As indicated below, the resistance to volume compression is at least an order of magnitude larger than the resistance to bilayer thickness and area changes.

The tight lateral packing of lipid molecules in the bilayer underlies its extremely low ion permeability and relatively low water permeability (85). However, this feature also contributes to the bilayer's resistance to area expansion. This is because even slight additional separation of lipid head groups (i.e., ~2%) will allow more water to enter between the acyl chains and destabilize the hydrophobic cohesive structure. Aspiration of spherical vesicles indicates a simple linear relation between membrane tension (t) and the relative area expansion of the bilayer

(1)

where A is the increase in surface area, A₀ is the original area, and K_A is the area expansion modulus (108). Typical values of K_A range between 10² and 10³ mN/m depending on the cholesterol content of the bilayer, while lytic tensions range between 3 and 30 mN/m, consistent with bilayers only being able to be expanded 2-4% before rupture (108, 296, 298). With the assumption of a bilayer compressibility modulus of 10⁹ N/m², the bilayer is at least 10-fold more compressible in area than in volume (given a K_A of 200 mN/m divided by 3 nm for the bilayer thickness). Thus, at near lytic tensions, the area may increase by ~4%, but the volume by <0.4% so that the thickness will decrease by 3.6%. Thus any fractional change in area should be accompanied by a proportional change in membrane thickness (h) so that

(2)

where h₀ is the unstressed membrane thickness and the expansion and thickness moduli are related according to K_A = K_h · h₀ (108). Estimates of K_h based on capacitance measurements of bilayer thickness changes during electrocompression indicate values of ~2 × 10⁷ N/m² (5), which would predict a K_A of ~70 mN/m assuming a bilayer membrane thickness of 3 nm (108).

Bilayer vesicles generated from membranes of RBC (183, 295, 296) and skeletal muscle (298) give values of K_A of around 500 mN/m, similar to the K_A measured for osmotically swollen (i.e, spherical) RBCs (106, 183). This agreement has been taken to indicate that the CSK of the RBC does not limit the elastic expansion of the bilayer (108). In studies of other cell types in which significantly lower values of K_A have been reported (e.g., 2-20 mN/m, e.g., see Ref. 187), it is not clear that elastic membrane expansion at constant area was being measured, since the cells were not preswollen to reduce excess membrane surface area. Interestingly, in the case of RBCs, it has been reported that K_A values vary by ±40% with voltage changes of ±200 mV (210, 211). Although the mechanism of this polarity-sensitive K_A effect remains unknown, it may reflect electric forces acting on packing of the highly asymmetrical lipid bilayer of the RBC (458). In this case, it will be interesting to examine the voltage effects on RBCs that have lost their phospholipid asymmetry due to lipid scrambling (152, 458).

The resistance of the bilayer to bending arises because of differential expansion or compression of the two monolayers within the bilayer and will depend on how tightly the two halves of the bilayer are coupled (i.e., degree of interdigitation between the acyl chains). If there is no coupling so that the two monolayers can rapidly slide past one another, there will be little resistance to bending. Bending rigidity is also dependent on the spontaneous curvature of the bilayer, which depends on the lipid composition and area of each monolayer (see sect. III) as well as the coupling of the CSK to the bilayer (91, 431). In the specific case of a bilayer sealed tightly to the walls of the patch pipette, the bending rigidity of the membrane patch should be increased because the attachment of the outer monolayer to the pipette walls will restrict its movement relative to the inner monolayer. Although the estimated bending modulus of the bilayer (K_B ~10¹⁹ N · m) indicates that bending resistance is significantly less than resistance to expansion (108), it is the bending rigidity that determines the shape of the lipid vesicle, its elastic response to membrane dimpling, and the amplitude of thermally induced fluctuations in the vesicle (452a; 276a and references therein).

Above the phase transition temperature, the bilayer has unrestricted internal fluidity and displays negligible surface shear rigidity so that it flows like a fluid in response to shear or extension. Below the phase transition, the shear rigidity increases along with hydrocarbon chain order (108). Furthermore, bilayers that undergo lipid phase separations may be expected to show heterogeneities in shear rigidity (138). Similarly, the existence of lipid rafts in cell membranes (369a) may result in differential rates of lipid flow in response to shear. However, more important is the cortical CSK that by providing the fluid bilayer with a solid support significantly increases the shear rigidity of the cell membrane and thereby allows elastic responses to membrane extension (see sect. VII). For example, the human RBC has an elastic shear modulus of ~10² mN/m and can recover rapidly from large extensions. However, after treatments {e.g., >48°C or increase in intracellular Ca²⁺ concentration ([Ca²⁺]_i)} that disrupt the cortical CSK, the shear modulus is so diminished the RBC undergoes spontaneous fragmentation (e.g., see Fig. 8 in Ref. 284; Ref. 152).

Elastic solids respond instantaneously to deformation. However, a bilayer vesicle cannot be deformed instantaneously because of the inertia of surrounding water movement and in specific cases possibly due to the viscous properties (e.g., bending viscosity) of the bilayer itself (24). For bilayer expansion, hydrodynamic drag rather than expansion viscosity is most likely rate limiting. For example, based on fluorescence polarization measurements of the bilayer hydrocarbon interior, the expansion viscosity (_A) of the bilayer has been estimated to be 10¹⁰ N · s/m (i.e., comparable to a 10-Å-thick layer of olive oil, Ref. 108). In this case, the time constant for area relaxation (_A) will be 10⁹ s according to the relation _A = _A/K_A and assuming a K_A of 10² mN/m (108). How does this value compare with experimentally measured kinetics of bilayer expansion? Clearly such measurements are limited by current methods. For example, even relatively sophisticated pressure-clamp techniques can only give pressure steps with a rise time of ~1 ms (272, 273), and these cause membrane patch movements (i.e., expansion) and MG channel activation with millisecond latencies (448). In comparison, voltage steps (i.e., with a rise time <10 µs) applied to outer hair cells cause membrane patch expansions of ~1% with a  of 100 µs, while the underlying membrane charge movement has a  of 10 µs (119a). The discrepancy may reflect membrane damping by the fluid movement in the pipette and/or the viscous drag of the cortical CSK. The relative contributions may be separated by measurements on blebbed (i.e., CSK-disrupted) hair cells. In terms of shear relaxation, the negligibly small shear modulus of the fluid bilayer should allow even faster intrinsic kinetics (i.e., <10⁹ s). The shear relaxation of cell membranes may be rate limited by the larger shear viscosity of the underlying CSK. In the specific case of bilayer bending, the interfacial drag between the two monolayers may be sufficiently large (i.e., 10⁷ N · s/m³) to slow membrane bending and shape recovery (110). Evidence of slow bending kinetics may be reflected in the relaxation of fine membrane tethers drawn from lipid vesicles (181) and possibly the adaptation of MG channel activity in liposome membrane patches (see sect. VIIIE).

In summary, the mechanical equilibrium of a lipid vesicle is established by the balance of external forces applied to the membrane (i.e., expansion and bending) opposed by the action of membrane tension and the bending rigidity. In considering deformation-sensitive membrane parameters that might influence membrane protein conformational changes, it is often overlooked that dilation of the elastic bilayer (i.e., increasing the area occupied by lipid molecules) should be accompanied by a proportional decrease in bilayer thickness (i.e., assuming volume incompressibility). In addition, and as described below, factors that affect spontaneous membrane curvature and bending rigidity may influence protein conformational changes. In terms of bilayer dynamics, that rate of membrane deformations involving bilayer expansion and extension may be damped by the hydrodynamics of the adjacent water compartments. However, membrane bending and relaxation may be rate limited by viscous drag between the monolayers.

	III. MECHANICAL DEFORMATION OF THE BILAYER BY MEMBRANE PROTEIN INSERTION

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The next level of complexity that can be considered is how insertion of membrane proteins may mechanically distort the bilayer and, in turn, how mechanically induced bilayer distortions may influence protein conformational changes. The central idea to bilayer-protein interaction is that the hydrophobic thickness of the bilayer immediately adjacent to the membrane protein will tend to match the length of the protein's hydrophobic exterior (Fig. 1; Refs. 290, 291). This may be expected to occur because any uncompensated mismatch will add a high energetic cost by exposing hydrophobic groups to water. Because proteins are relatively rigid, whereas lipid hydrocarbon chains are flexible, the condition of hydrophobic matching can be achieved by stretching, squashing, and/or tilting of the lipid chains (172, 193). Recent direct evidence supporting protein-induced changes in lipid organization comes from the demonstration that hydrophobic -helical peptides, including gramicidin A, can change a bilayer into a nonlamellar structure, with this transition dependent on the degree of hydrophobic mismatch (217). Furthermore, insertion of peptides, including alamethicin, into bilayers causes a concentration-dependent thinning of the bilayer as measured by lamellar X-ray diffraction (173).

View larger version (34K): [in this window] [in a new window]   Fig. 1. A membrane protein in changing conformation also undergoes changes in hydrophobic mismatch with the lipid bilayer. A: positive hydrophobic mismatch in which the protein promotes local positive curvature in the lipid bilayer. B: a protein conformation that promotes local negative curvature. C: neutral mismatch in which the protein does not distort the bilayer. [Modified from Fattal and Ben-Shaul (111).]

One of the consequences of the hydrophobic mismatch idea is that proteins will tend to surround themselves with lipids of matching size and shape so that the mechanical strain on the bilayer will be minimized (82, 111, 125, 140, 352). Furthermore, if the lipid composition of the membrane can be altered, one might expect to see shifts in protein conformational changes. Evidence supporting these ideas has come from studies of the effects of foreign phospholipids and lipophilic agents on MG channel activities (53, 251, 253). For example, the opposite effects observed with some lipophilic agents on gramicidin and N-methyl-D-aspartate (NMDA) channel kinetics may be explained by differences in lipid shape, as defined by the ratio of head group size (H) to the acyl tail area (T), on the localized curvature of the bilayer (53, 253). Lipids with H = T will tend to favor neutral curvature, lipids with H > T will favor positive curvature, and lipids with T > H will favor negative curvature. As described below, the effects of lipids of different geometry on MG channel gating provided the initial clue that changes in membrane thickness and/or local curvature may underlie one mechanism of MG channel gating (136, 251). The importance of the surrounding lipids on protein function may also be reflected in the enzyme-regulated (i.e., floppase and translocase) asymmetrical distribution of phospholipid in each monolayer, which is lost with scramblase activation (458).

Figure 1 considers the specific case of a membrane protein that has three stable conformational states, each with different types of hydrophobic mismatch with the bilayer (i.e., A positive, B negative, and C neutral). Insertion of foreign lipids or membrane thinning by altering the energetic cost of membrane deformation should cause shifts in the distribution among these conformations (e.g., membrane thinning would favor B). However, effects such as ligand binding, phosphorylation, or membrane polarization may cause shifts independent of hydrophobic mismatch. In this case, complex interactions may arise between mechanical and other forms of stimuli.

	IV. SIMPLE PEPTIDES THAT FORM MECHANICALLY GATED CHANNELS

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Alamethicin and gramicidin are the best-characterized membrane channels in terms of their biophysics (i.e, conductance and gating), structure-activity relations, and modeling of open-closed channel conformations (6, 47, 355, 426). Therefore, the demonstration that both channels display mechanosensitivity in pure lipid bilayers, similar to prokaryotic MG channels, has provided the opportunity to analyze possible underlying molecular mechanisms in extremely well-defined simple systems.

Alamethicin is a 20-amino acid peptide that forms voltage-gated multi-conductance state channels in lipid bilayers (47, 355). The most commonly evoked model to explain channel formation is a "barrel-stave" model in which each stave of the barrel is formed by a single -helical monomer. To explain the voltage sensitivity of alamethicin, two main classes of mechanisms have been proposed (47). In one, the channel exists as a preaggregate of subunits, and a voltage-dependent conformational change results in channel formation. In the other, alamethicin exists predominantly as monomers and voltage-dependent insertion (or partitioning) with subsequent aggregation that leads to channel formation. Because evidence exists supporting both mechanisms (47), analysis of the mechanosensitivity of alamethicin channel gating was used in an attempt to discriminate between the two mechanisms (313). Initially, Opsahl and Webb (313) demonstrated in patch-clamped bilayers (i.e., using the "tip-dip" method) that increased membrane tension increased the probability of the channel occupying higher conductance states. Their quantitative analysis of the relation between the state of occupation and applied membrane tension (t) was interpreted in terms of the work done in channel opening W = t · A, where the switching between the adjacent states involves an increase in membrane occupied area of the channel complex (A) of ~1.2 ± 0.10 nm². Based on these area changes, they proposed that the mechanosensitivity of switching between different conductance states could be explained by a model involving two tension-sensitive steps. Step 1 involved insertion or partitioning of an additional monomer (cross-sectional area ~0.8 nm²) into the channel bilayer. Step 2 involved the subsequent association of the inserted monomer with the existing channel aggregate resulting in an increase in pore area (~0.4 nm²). In favoring the subunit-recruitment model, Opsahl and Webb (313) pointed out that the observed area changes were most likely too large to be compatible with a model involving rearrangement (expansion) of the monomers within a fixed aggregate (115a). Furthermore, their observation that the free energy difference between closed and open channel conformations varied linearly with tension confirmed that first-order area changes were responsible for the mechanosensitivity, while second-order (quadratic) effects due to compliance changes in channel states were not significant (69, 349). An increase in pore area of 0.4 nm² would give an increase in pore volume of ~1.3 nm³ assuming a pore length of 3 nm. This volume change is at least of the same order predicted based on osmotic experiments that indicate channel switching involves uptake of up to 3 nm³ of water (421). Opsahl and Webb (313) demonstrated equivalent tension sensitivity for the three lowest adjacent conductance states in bilayers of fixed composition. It will be interesting to see if these same states as well as the higher states generated in bilayers with high curvature (214) display the same tension sensitivity as might be expected from their model. Finally, Opsahl and Webb (313) did not consider the free energy contribution on changing the environment of one face of the recruited alamethicin subunit from that low dielectric of the lipid bilayer to the high dielectric of the aqueous pore. As described later for a bacterial MG channel, the large free energy change associated with channel opening may arise from the energetic cost of exposing pore-lining, hydrophobic residues to water in the open pore (443, see sect. VII).

For alamethicin, the subunit recruitment model remains intuitively attractive and appears consistent with most of the experimental data. However, data on the effects of lipid composition on alamethicin conductance switching (214) may indicate an alternative mechanism related to hydrophobic coupling between the peptide and bilayer.

Gramicidin is a 15-amino acid peptide that also forms channels in bilayers. Although a simple molecule, it exhibits two different folding motifs: a double helix and a helical dimer. This polymorphism in structure is manifest in solution, in bilayers, and in the solid state (426). Although both folding motifs may form transmembrane pores or channels, there is substantial evidence that the most frequently observed channel arises through the dimerization reaction between two nonconducting monomers that insert into each monolayer as -helices (306). The length of the gramicidin dimer exterior is 2.2 nm, which can be compared with ~3 nm for a phopholipid bilayer and 3.2 nm for the long axis of alamethicin (115a). As illustrated in Figure 1B, because of gramicidin's negative mismatch, the bilayer hydrophobic core will tend to be compressed (i.e., seen as a negative curvature) to match the channel's hydrophobic exterior surface.

Several groups have previously reported apparent tension sensitivity in the gating of gramicidin channels (101, 297, 341). However, interpretation of these early studies was complicated because channels were studied in black lipid bilayers where changes in tension remain undefined and in some cases membrane thickness as well as tension was altered. In a more recent study, the pipette aspiration technique was used to increase tension in bilayer vesicles (136, see sect. II). With this technique it was demonstrated that tension increased the rate of gramicidin channel formation (2- to 4-fold) and, to a lesser extent, the average channel lifetime (136). Note this increase in lifetime was opposite to previous reports that increased tension decreased gramicidin channel lifetime (101, 297, 341).

For gramicidin, there is little difference in the membrane area occupied by monomers and dimers that might explain the tension sensitivity of channel gating. Instead, Goulian et al. (136) proposed that increased tension, by causing bilayer thinning (see sect. IIB), improved the hydrophobic coupling between the bilayer and the dimer, thereby reducing the membrane deformation energy associated with channel formation and increasing the activation energy associated with channel dissociation. The smaller effect of tension on rates of channel dissociation compared with rates of formation were shown to be consistent with the smaller displacement (~0.1 nm) of monomers necessary for dissociation compared with the larger mismatch (i.e., ~0.4 nm) between the dimer and bilayer. In contrast to the linear dependence of free energy change of alamethicin channel switching with tension (313), gramicidin displayed a quadratic dependence (136), possibly reflecting the elastic properties of membrane deformation (252). According to Nielsen et al. (300), if a protein conformational change involves an increase in the hydrophobic mismatch from 0.1 to 0.13 nm, there will be a 10-fold shift in the equilibrium distribution of protein conformations due to these elastic properties of the membrane.

In addition to the effects of membrane tension, a variety of other experimental maneuvers have been shown to alter gramicidin channel gating. These effects were also interpreted as arising through changes in membrane deformation energy. The treatments include incorporation of membrane lipids, detergents, and cholesterol that tend to alter the membrane local curvature, thereby either stabilizing the channel, in the case of agents that promote positive curvature, or destabilizing the channel, in the case of agents that promote negative curvature (251, 253). Similarly, conditions such as elevated Ca²⁺ that decrease electrostatic energy of the bilayer by screening surface charge are proposed to promote channel dissociation by a mechanism involving changes in the membrane local curvature or thickness (254). Note that although the gramicidin channel is axially symmetrical, its sensitivity to the sign of local curvature of the adjacent lipid is expected in terms of the mismatch model (see Fig. 1). One would also predict that changes in membrane voltage, by altering membrane thickness through electroconstriction (5), would also alter membrane deformation energy and thereby gramicidin channel gating. However, voltage-sensitive gating of gramicidin has not been reported (6).

In summary, increased bilayer tension promotes dimerization of gramicidin and higher conductance states of alamethicin. It may be that different mechanisms underlie the mechanosensitivity of the two channels. For alamethicin, a subunit recruitment model has been favored over the fixed aggregate model. However, there is evidence that indicates the switching between alamethicin conductance states is dependent on lipid composition, similar to gramicidin except that the lipids that promote gramicidin dimerization favor lower conductance states of alamethicin (214, 251). Because other evidence indicates that membrane deformation energy may be the major driving force for the alamethicin insertion transition (see Ref. 171), the hydrophobic mismatch model may also contribute to the mechanosensitivity of alamethicin. Finally, the existence of polymorphic structures, ambiguities, and unresolved issues with such simple channel-forming peptides is a useful reminder of the complications that lie ahead for modeling larger and more complicated channel proteins as described in the next section.

	V. STRUCTURE OF PROKARYOTIC CELLS

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The discovery of Archaea (formerly achaebacteria) has clearly shown that the old prokaryote/eukaryote dichotomy is obsolete by largely oversimplifying diversity of prokaryotic microbes in all its aspects including prokaryotic cell envelopes (436). Nevertheless, from the perspective of the MG channel gating mechanism, bacterial as well as archaeal cells may be considered the next level of complexity after a bilayer vesicle. The cytoplasmic membrane of bacteria is a fragile structure composed of phospholipids and proteins enclosed by a cell wall (i.e., outer membrane) that provides a strong, rigid structural component able to withstand the osmotic pressures caused by the intracellular concentration of various osmoticants in the cell (423). Without the mechanical support of the cell wall, a bacterial cell would behave as a tiny dialysis bag that would take up water from the environment, swell, and burst. Bacterial cell walls (with the exception of the mycoplasma) have a structural component called peptidoglycan that provides the rigidity necessary to maintain cell integrity. The major building blocks of peptidoglycan are N-acetylglucosamine and N-acetylmuramic acid that are unique to bacterial cells. On the basis of the Gram stain, a differential staining technique invented in 1884 by Christian Gram, bacteria can be divided into two large groups: Gram positive and Gram negative, which differ in the peptidoglycan content of their cell wall (i.e., ~5 times larger in Gram-positive cells) (Fig. 2A) than in Gram-negative cells (see Fig. 2B). In addition, Gram-negative bacteria have a second chemically distinct outer membrane attached to the peptidoglycan layer on its external side. The lipid bilayer of the outer membrane is made of phospholipids in the inner leaflet of the bilayer and lipopolysaccharides (LPS) in its outer monolayer. Gram-positive cells lack the second outer membrane (300b).

View larger version (37K): [in this window] [in a new window]   Fig. 2. Cell envelope of Gram-positive (A) and Gram-negative bacteria (B). [From Volk (423).]

The situation with Archaea is more complicated, likely reflecting the large diversity of extreme habitats to which Archaea have adapted (318). Archaea lack peptidoglycan, which was one of the features used originally to define prokaryotes. Two types of archaeal cells, which to date were found to harbor MG channels in their cell membranes, may illustrate this. The cell wall in halophilic archaea, such as Haloferax volcanii, whose cell membrane was the first to be examined for the presence of MG channels (239), consists of the S layer formed by a hexagonal arrangement of a glycoprotein (238). The protein appears to be anchored in the cytoplasmic membrane by a hydrophobic stretch found near the COOH terminus of the H. volcanii glycoprotein sequence (392). Thermoplasma volcanii is the second archaeon found to have MG channels in its cell membrane (220). This thermophilic archaeon has no cell wall, but instead contains an outer meshlike lattice of elements similar to nuclear lamins that is reminiscent of the CSK in animal cells (179). In addition, its cell membrane contains ether lipids based on 40-carbon, isopranoid-branched diglycerol tethraethers (361). In general, lipid bilayers of cell membranes of all archaea consist of diphytanylglycerol-diether or -tetraether or both (92). It is worth mentioning that neither bacteria nor archaea have a CSK in the eukaryotic sense. To describe the structural diversity of prokaryotic cell envelopes in its entirety would go over the scope of this review. What matters from the perspective of prokaryotic MG channels is that despite this diversity, it is always the lipid bilayer that is the tension-bearing element. The cell wall functions as a parallel viscoelastic structure that constrains the bilayer from excessive dilation and in this way may reduce MG channel activation (41, 265, 267). The practical implication of the MG channel's sensitivity to bilayer tension is that, as discussed later, they retain their mechanosensitivity when reconstituted in liposomes (239, 388). The physiological implication is that they regulate cellular turgor by responding to bilayer stretch caused by osmotic swelling (2, 244; see sect. VIJ).

	VI. MECHANICALLY GATED CHANNELS IN BACTERIA AND ARCHAEA

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Since their initial discovery in giant spheroplasts of E. coli (267), the existence of MG channels in Gram-negative and Gram-positive bacteria has been amply documented (17, 18, 26, 27, 268, 387, 455, 456). The best-characterized are the MG channels of the Gram-negative bacterium E. coli, which have been studied by the patch-clamp technique in various giant spheroplasts (41, 76, 229, 266, 267) and in reconstituted membrane fractions fused with liposomes (18, 87). Based on their conductance and sensitivity to applied pressure, three types of mechanosensitive channels (Msc) can be distinguished: MscM (M for mini), MscS (S for small), and MscL (L for large) (17). The higher the conductance, the higher their activation pressure as evident in both in situ or in vitro recordings. Also, in Gram-positive S. faecalis and B. subtilis (397, 398, 455), membrane stretch results in the activation of a whole array of conductances, ranging from 100 pS to up to several nS.

The property of E. coli MG channels of being activated by mechanical force transmitted via the lipid bilayer (266) allowed for detergent solubilization and functional reconstitution of these channels into artificial liposomes amenable to patch clamp (388). Furthermore, it allowed application of a unique strategy of fractionation of E. coli membrane constituents by column chromatography and functional examination of the individual fractions for MG channel activity by patch clamp. This unusual approach led to the identification of a membrane protein underlying the activity of MscL (386, 388). The MscL protein was partially sequenced, which enabled the cloning of the corresponding mscL gene (384). The expression of the mscL gene alone in a heterologous and in an in vitro transcription/translation system demonstrated that the mscL gene alone was necessary and sufficient for MscL activity.

The mscL gene encodes a 15-kDa protein comprising 136 amino acid residues corresponding roughly to the 17-kDa protein band on a SDS-PAGE originally identified as the MscL protein (386, 388). Manipulation of the mscL gene by recombinant DNA techniques enabled production of the MscL protein on a preparative scale, which in addition to functional studies allowed also for higher order structural studies of MscL. Two methods have been used for simple purification of the MscL recombinant proteins. The first method employs the glutathione S-transferase (GST) protein fusion technique to express MscL attached to a cleavable GST domain (168), whereas the second method uses a 6xHis polyhistidine tag for purification of the recombinant MscL protein on a Ni⁺-NTA column (27). Both methods yielded functional MG channels in patch-clamp experiments. The 6xHis-tagged MscL protein was used for secondary structure analysis by employing both transmission Fourier transform infrared spectroscopy (FTIR) and circular dichroism (CD) (7). The MscL secondary structure includes two -helical transmembrane domains (M1 and M2) connected by a periplasmic loop (Fig. 3, A and B). Thus MscL belongs to the family of structurally related ion channels with two membrane segments that include the epithelial sodium channel (ENaC), the inward-rectifier potassium channel (Kir), and the ATP-gated (P2X) cation channel (37, 302). Using the PhoA fusion technique, Blount et al. (27) could demonstrate that the NH₂ as well as the COOH terminus of MscL were located within the cytoplasm.

View larger version (32K): [in this window] [in a new window]   Fig. 3. Large mechanosensitive channel (MscL) membrane-spanning models. A: membrane topology of E. coli MscL monomer proposed by Sukharev et al. (387). B: drawing of the MscL pentamer (left) and monomer (right) from M. tuberculosis according to the 3-dimensional structural model proposed by Chang et al. (60). [Modified from Oakley et al. (305).]

The higher order structure of MscL was first assessed in cross-linking studies. In one study, the multimeric structure of MscL was examined either by cross-linking the protein in situ and visualizing the cross-linked products by Western blotting using MscL antibodies against a COOH-terminal peptide or by cross-linking purified 6xHis-tagged MscL in which purified proteins were cross-linked. Both approaches indicated that MscL might form homohexameric channels (26, 27). In another study, various cross-linkers were applied directly to bacterial cells in which the MscL protein was radiolabeled by [³⁵S]methionine (170) and left open the possibility that MscL forms multimers of higher order other than a hexamer. However, recent reevaluation of cross-linking experiments using various cross-linking reagents indicates that MscL is a homopentamer (389, 270a). Preparative scale production of milligram amounts of the MscL protein allowed employment of electron crystallography to study the structural assembly of MscL. The crystallographic analysis of two-dimensional MscL crystals at 1.5-nm resolution indicated that MscL forms homohexameric channels in lipid bilayers (353). However, the three-dimensional X-ray crystallographic study by Chang et al. (60) solved the oligomeric structure of the MscL homolog from Mycobacterium tuberculosis (Tb-MscL) to 0.35-nm resolution and indicated this homolog is organized as a homopentamer (Fig. 3C) (for discussion of homohexamer versus homopentamer, see sect. VID).

MscL forms nonselective ion channels of a very large conductance. The absence of any cation/anion selectivity or saturation in channels conductance up to 2 M KCl indicates MscL forms a large water-filled pore (75, 390). The reported values for MscL conductance range from 0.9 to 4.4 nS. Some of the differences may reflect the different ionic composition of the recording solutions used for patch-clamp experiments. However, the variations in MscL conductance were also observed in recording solutions of the same or similar ionic composition (27, 75, 226, 283, 384). The two extreme conductance values reported for MscL (in a recording solution containing 200 mM KCl plus 40 mM MgCl₂ and 5 or 10 mM HEPES, pH 6.0 or 7.0) are 2.5 and 3.8 nS, although the exact reason for these diffferences remains unclear. As discussed in the next section, one explanation is that MscL can form more that one type of homomultimeric channel, perhaps analogous to the channel and pore forms of gramicidin (see Ref. 426).

It is important to establish the multimeric organization of MscL because, as previously discussed for alamethicin and gramicidin, this feature can have important implications for gating mechanisms. As indicated above, homohexameric (Eco-MscL) and homopentameric (Tb-MscL) structures have been reported from crystallographic analysis (60, 353). In addition, several reports based on cross-linking experiments contributed to the MscL structural controversy by showing that MscL may form hexamers (27) as well as pentamers (389). Moreover, tandems of two MscL monomers expressed as a single dimer protein formed functional channels in giant E. coli spheroplasts, indicating that the functional channels can be made from an even number of monomers (27). On this last point, it would be interesting to obtain two- or three-dimensional crystals of channels made of the MscL protein dimers.

Is MscL a hexamer or a pentamer? Probably it is safe to say that MscL forms pentameric channels taking into account the resolution of 0.35 nm at which the Tb-MscL three-dimensional structure was obtained (60). Moreover, at the resolution of 1.5 nm for the two-dimensional Eco-MscL crystals, a pentameric MscL structure is just as likely to be judged as a hexamer (353). A resolution of at least 1.0 nm would be required to conclusively distinguish between hexameric and pentameric structures of the two-dimensional MscL crystals (J.-L. Rigaud and J-.J. Lacaperre, personal communication). Nonetheless, it is worthwhile pointing out several peculiar details of the Tb-MscL structure and the experimental conditions at which the three-dimensional structure was obtained. According to the Protein Data Base (PDB) summary report, the Matthews coefficient (V_m) of 5.98 for MscL is high compared with known structures of other proteins in the data base, indicating that the MscL structure is an outlier. The coefficient is usually in the range between 1.5 and 4.0 for tightly and loosely packed proteins, respectively. Also, the Ramachandran Z-score of 5.671 for MscL appears very low and suggests a very unusual backbone conformation of Tb-MscL that is probably due to some local uncertainties in the backbone side chain conformation (A. Oakley, personal communication). In addition, the Tb-MscL crystallization experiments were performed at a very low pH between 3.6 and 3.8. This pH is below the pK_a value of 4.25 of the glutamic acid residue E104 in the COOH-terminal domain of Tb-MscL, which probably had to be protonated to stabilize the MscL pentameric structure. At higher pH one might expect the five E104 residues of the pentamer to become negatively charged causing destabilization of the multimeric structure, unless there is charge compensation by neighboring basic residues or cations present in the surrounding medium (305). Change in pH is known to modulate significantly the pressure sensitivity of MG channels (as discussed later) and also induce structural changes in proteins (42). Therefore, at pH ~7 the MscL tertiary structure might differ from the reported pentameric one.

Obviously, it is not possible to compare directly the two- or three-dimensional structural data of MscL with the electrophysiological conductance measurements, since the crystallographic structures are depicting closed channels, whereas electrophysiology can detect only open conducting channels. For example, the three-dimensional crystal structures of the Tb-MscL pentamer revealed a transmembrane pore that narrows from 3.6 to 0.4 nm in diameter in going from the extracellular to the cytoplasmic surface (60) (Fig. 3B). In contrast, patch-clamp permeation studies indicated the open MscL has a channel pore of ~4.0 nm diameter (75). Consequently, a major structural rearrangement occurs when MscL "switches" between the two conformations (14). To fully understand the process will most likely require the crystal structure of the open MscL.

Bacterial and archaeal MG channels provide a clear demonstration that microbial MG channels can sense membrane tension directly. The tension develops in the lipid bilayer alone and directly gates these channels as described by the bilayer model. Since it was first proposed in relation to bacterial MG channels (261, 266), the bilayer model has become, along with the tethered model, one of the two mechanisms used to describe MG channel gating (see sect. VIIIC). In the case of MscL, the validity of the bilayer model has been amply documented (28, 168, 384, 387, 388, 390). Interestingly, the well-studied stretch-activated cation (SA-CAT) channel endogenous to Xenopus oocytes also appears to be gated by bilayer tension (448).

As the amount of negative pressure (i.e., suction) applied to a patch pipette increases, the MscL channel open probability also increases (Fig. 4A). Activation of MscL by pressure can be described by a Boltzmann distribution function for the channel open probability (P_o) (Fig. 4B)

(3)

where p is the applied negative pressure, p_1/2 is the suction at which the channel is open half the time, and  is the slope of the plot ln [P_o/(1  P_o)]. Because the Boltzmann function is very often used to characterize the mechanosensitivity of MG channels in the literature, we will discuss the meaning of the terms p_1/2 and , first in relation to MscL and then other MG channels. The following exercise should be useful to laboratories studying MG channels that do not routinely image membrane patch movements and calculate the membrane tension changes.

View larger version (15K): [in this window] [in a new window]   Fig. 4. Activation of MscL by negative pressure applied to the patch-clamp pipette. A: current traces of the reconstituted recombinant 6xHis-MscL recorded from an isolated liposome patch at various negative pressures applied to a patch pipette. Trace is a ~100-s-long recording at a pipette potential of +30 mV. C denotes the closed state, and On denotes the open state of n number of channels. Nine channels were active in the patch shown. B: curve is a Boltzmann distribution function calculated for a single MscL shown in A. The function relates negative pressure (suction) and open channel probability (Po). Boltzmann distribution for a single MS channel has a form Po/(1  Po) = exp [(p  p1/2], where p is the applied negative pressure, p1/2 is the suction at which the channel is open 50% of the time (Po = 0.5), and  is the slope of the plot ln [Po/(1  Po)]. The Boltzmann function is very often described in the MS channel literature in terms of p1/2 and . For the 6xHis-MscL in the particular recording shown in A, p1/2 was estimated to be ~77 mmHg, whereas the sensitivity to pressure 1/ was ~5 mmHg for e-fold change in Po, which corresponds to Go  15kT (Eq. 5). Overall, Go for the recombinant 6xHis-MscL was estimated to be 17.0 ± 1.5kT (SE; n = 4) (A. Kloda and B. Martinac, unpublished data).

As shown previously (147, 374, 375, 390, 448), most MG channels respond to mechanical forces along the plane of the cell membrane (membrane tension), and not pressure perpendicular to it. According to the model of Howard et al. (190), the free energy (G) is a linear function of membrane tension t

(4)

where G_o is the difference in free energy between the closed and open conformations of the channel in the absence of the externally applied membrane tension and A is the assumed difference in membrane area occupied by an open and closed channel at a given membrane tension, whereas tA is the work required to keep an MG channel open by external mechanical force at the open probability of 0 < P_o < 1. Consequently, in this model, the size of A is considered the sole parameter determining the mechanosenitivity of channel gating (i.e., the larger A the more sensitive the channel). Evidence indicates that for non-MG channels, like the ACh receptor channel (417) and specific voltage-gated K⁺ channels (56, 130), the movement of transmembrane helices is quite small. In contrast, large movements of MscL are needed to account for the large pore formation and the steep tension-response relation of MscL (390).

Using the expression in Equation 4, the Boltzmann function can be rewritten as

(5)

In the case of MscL, which is activated by membrane tension near the lytic strength of the bilayer (i.e., maximal curvature), it has been demonstrated that in that activating pressure range and assuming an elastic membrane with a K_A = 10² mN/m (see Refs. 373, 390), the tension will be nearly proportional to the pressure (i.e., there will be little change in curvature). Using a version of Laplace's law

(6)

where r is the radius of curvature of the liposome membrane patch under external negative pressure p applied to the patch pipette. Thus, under these conditions, p_1/2 and t_1/2 , the characteristic negative pressure and membrane tension, respectively, at which the channel is open 50% of the time can be assumed to remain constant for a membrane patch during an experiment. Because the free energy difference G (Eq. 4) is equal to zero when the open probability P_o = 0.5 (P = p_1/2 and t = t_1/2), consequently t_1/2 = G₀/A and p_1/2 = 2G₀/rA, whereas  = rA/2kT. Thus it follows that at least in liposome or bleb experiments in which parallel CSK elements are not variables in supporting bilayer tension, the slope term will provide a direct estimate of molecular rearrangements of a MG channel, as long as the diameter and shape of patch pipettes remain nearly constant throughout the experiments. A convenient expression can be obtained by multiplying p_1/2 and  

(7)

showing that the product _MGC is independent of the patch geometry. _MGC provides a direct estimate of the energy difference G_o between the closed and open state of an MS channel, and thus it presents the very characteristic of any type of MG channel reconstituted into a defined lipid membrane. For example, according to several reports, a negative pressure (p_1/2) ranging between ~7.7 and 10.3 kN/m² (i.e., ~58 and 77 mmHg) is required to activate MscL 50% of the time in liposome patches (3, 141, 246a, 220b), whereas the sensitivity to pressure (1/) of the MscL channels was found to vary between 0.6 and 0.8 kN/m² (i.e., 4.5 and 6.0 mmHg) with  ~1.67 and 1.25 (kN/m²)¹ (i.e., 0.22 and 0.17 mmHg¹), respectively (3, 75, 168, 220b). It follows that on average _MscL ~14, and consequently, the average energy required for opening MscL is G_o ~14kT. This value is in reasonable agreement with an independent estimate of ~18.6kT obtained for G_o from P_o-membrane tension curves for MscL (390). Clearly, the interpretation of Equation 7 becomes more complicated for cell membranes in which the associated cortical CSK (i.e., an extrinsic factor) can alter the tension seen by the bilayer and thereby change the shape of the Boltzmann independent of the intrinsic properties of the channel protein (see Ref. 447 and sect. VII).

1) Lysozyme, which disrupts the peptidoglycan of the bacterial cell wall, irreversibly increases the pressure sensitivity of MG channels in E. coli giant spheroplasts (41, 268).

2) Cytochalasin, which disrupts actin microfilaments, increases the pressure sensitivity of MG channels in chick muscle and snail neurons (142, 370).

3) Mechanical decoupling of the CSK from the plasma membrane (i.e., blebbing) in Xenopus oocytes decreases the MG channel mechanosensitivity (154, 160, 447).

4) Amphipatic (amphiphilic) compounds can increase mechanosensitivity of bacterial (266) and eukaryotic MG channels (323, 372).

5) Deletion, substitution, or single-site mutations can either increase or decrease the native MscL mechanosensitivity (25, 28, 170, 316, 317).

6) Changes in pH can affect MG channel sensitivity in both ways, with alkaline pH increasing (143; Martinac, unpublished data) and acidic pH decreasing the channel's pressure sensitivity (28; Martinac, unpublished data).

These examples illustrate how the apparent mechanosensitivity of a channel may be altered by extrinsic, intrinsic, or possibly by a combination of mechanisms. Specifically, examples 1-4 may involve alteration in the way mechanical force is delivered to the channel (i.e., by alteration of CSK or bilayer properties) without altering the intrinsic properties of the channel protein (see also adaptation, sect. VIIIE), example 5 may involve changes in the protein's intrinsic mechanosensitivity, and example 6 may result from changes in both extrinsic (bilayer) and intrinsic (protein) properties.

In the specific case of MscL reconstituted into liposomes, the following example illustrates how, for a MG channel activated by lipid bilayer tension, the measured changes in p_1/2 and  can provide an estimate of the channel's intrinsic physical properties and identify the contribution of specific structural domains to the gating mechanism. Specifically, Ajouz et al. (3) studied the effects of various proteases on pressure-dependent gating of MscL in an attempt to identify molecular domains of MscL responsible for mechanosensitivity. They demonstrated that both parameters,  and p_1/2, were dramatically affected by protease treatment, with the slope term  significantly increased and p_1/2 decreased (i.e., mechanosensitivity increased). The quantitative changes in p_1/2 and  caused by protease treatment were such that _MscL (Eq. 7) ranged between 12 and 18 (i.e., G₀ ~12-18kT) compared with ~14 before protease treatment. Because _MscL and G₀ were hardly affected by cutting the extramembraneous domains, it was concluded that neither the cytoplasmic termini (i.e., the NH₂ and COOH termini) nor the S2-S3 periplasmic loop (see Fig. 3A) contribute critically to the mechanical gating of MscL. Instead, it was proposed that these extramembranous domains resist the movement of the transmembrane helices that underlie the critical event in mechanical gating of MscL. Note this result disagrees with the electromechanical model described below that proposes the NH₂ termini gate MscL (141).

Based on a mutagenesis study in which Gly-22 (E. coli MscL) was changed to all other 19 amino acid residues, Yoshimura et al. (443) concluded that by analogy with Ala-20 of Tb-MscL, Gly-22 should surround the Eco-MscL gate.

Indeed, the Tb-MscL has been found to be extremely stiff when expressed in E. coli and examined by the patch clamp. Specifically, Moe et al. (283a) found that the channel required twice the membrane tension needed to gate the Eco-MscL. In addition, their study showed that amino acid substitutions at the neighboring residue V21 had severe effects on the channel mechanosensitivity, indicating that besides G22, V21 also participates in the energy barrier of MscL gating (283a). By using the program VOIDOO/FLOOD (222), Oakley et al. (305) estimated that in the closed conformation MscL could be filled with water molecules to a point 2.4 nm below the periplasmic surface of the channel. Also, the second cavity at the cytoplasmic face of the channel was found to be water accessible. Consequently, most of the inside of the closed MscL contains water except for a hydophobic stretch of 0.8 nm that includes the Gly-22 residue and forms a water-tight occlusion. Thus the hydrophobic channel gate is quite thin compared with the 3- to 3.5-nm-thick membrane bilayer (60). If we accept that in the open state of MscL, the hydrophobic gate becomes exposed to water, as originally proposed by Cruickshank et al. (75) and later reiterated by Yoshimura et al. (443), then can the value of 18.6kT forG₀ be explained solely by this process. Taking into account that 17 mJ/m² is necessary to transfer a hydrophobic protein from an organic solvent into an aqueous environment (66), it follows that 18.6kT suffices to expose a hydrophobic area of 4.42 nm² to water (18.6kT = 7.521 × 10²⁰ J). This area is relatively small compared with the total membrane-associated area of MscL (~140 nm²) and roughly corresponds to a half surface of 5 -helices, each having a diameter 2r = 0.68 nm (396), with a height l = 0.8 nm corresponding to the height of the hydrophobic gate (i.e., 2rl ~5/2 = 4.27 nm²). Consequently, this result indicates that most, if not all, of the channel opening energy of 18.6kT is used to expose a relatively small buried hydrophobic surface