Method of Determining the State of Activation of a Protein

The present invention relates to a method for determining the state of activation of a (poly)peptide, comprising the steps of: (a) immobilizing a (poly)peptide on a carrier; (b) attaching the (poly)peptide immobilized on the carrier to a force measuring device of a force spectroscope; (c) applying a pulling force to the (poly)peptide and measuring the force required for stretching and/or unfolding of the (poly)peptide attached to the carrier, wherein the measurements are carried out (i) prior to and (ii) after treatment with a known modulator of said (poly)peptide; (d) comparing the force spectra of the (poly)peptide prior to and after treatment with the modulator; and (e) concluding from a difference of the force spectra of step (d) on the state of activation of the (poly)peptide. Furthermore, the present invention relates to a method for determining whether a compound is an activator of (poly)peptide function, comprising the steps of (a) immobilizing a (poly)peptide on a carrier; (b) attaching the (poly)peptide to a force measuring device of a force spectroscope; (c) applying a pulling force to the (poly)peptide and measuring the force required for stretching and/or unfolding of the (poly)peptide attached to the carrier, wherein separate measurements are carried out in the presence of a test and a reference buffer, wherein the test buffer comprises a compound suspected of being an activator of the function of said (poly)peptide and wherein the reference buffer comprises (i) a compound known to have no effect on the activity of said (poly)peptide or (ii) a compound known to be an activator of said (poly)peptide; (d) comparing the force spectra of the (poly)peptide measured in the presence of the test and the reference buffer; and (e) concluding from a difference of the force spectra of step (d) whether the compound suspected of being an activator of said (poly)peptide is an activator of the (poly)peptide. Finally the present invention relates to a method for determining whether a compound is an inhibitor of (poly)peptide function.

Several documents are cited throughout the text of this specification. The disclosure content of the documents cited herein (including any manufacture's specifications, instructions, etc.) is herewith incorporated by reference.

For apparent reasons it is of prime interest for the pharmaceutical industry to be able to study in detail the state of activation of proteins which are the targets of their drugs. Atomic force-microscopy (AFM) has recently been used to gain some insight into the molecular response upon treatment of the target proteins with specific drugs. However, these experiments did not allow to detect and to locate the molecular interactions activating these proteins. Since more than a decade the tip of the AFM cantilever has served as a nano-tweezer, enabling to manipulate biological objects at the molecular scale [29,38,57,71,111]. The outstanding positioning precision (≈0.1 nm) and force sensitivity (≈5 pN) of the AFM has made even the most delicate single-molecule unfolding experiments using force spectroscopy possible. In these experiments, the force applied to a single protein plays the role of a denaturant leading to complete unfolding of its three-dimensional structure. In their initial studies, Rief and co-workers applied single-molecule force spectroscopy to the giant muscle protein titin, which consists of repeats of globular immunoglobulin and fibronectin domains [111,113]. The continuous extension of the protein resulted in the subsequent unfolding of the globular domains allowing the unfolding force and pathway of each domain to be detected [79,113,149].

In contrast to many experiments performed on water-soluble proteins, the application of single-molecule force spectroscopy to membrane proteins [93,103] yielded surprisingly detailed insights into the inter- and intramolecular interactions stabilizing their three-dimensional structure. This has been demonstrated on membrane proteins like BR [59,98], halorhodopsin [18] from Halobacterium salinarum, human aquaporin-1 [85], and the Na⁺/H⁺ antiporter NhaA from Escherichia coli [64]. To select a membrane protein for a force spectroscopy experiment the protein containing membrane was first imaged at sub-nanometer resolution. Then the AFM tip and the selected protein are brought into contact. Applying a force of 300-1000 pN results in binding of one terminal end to the tip either via a covalent bond [103] or enforced non-specific adsorption [98]. Withdrawing the tip from the membrane stretches the terminus of the protein and causes the cantilever to deflect. Upon further separating the tip and surface, the force pulling on the protein steadily increases. As soon as the force exceeds the stability of the protein it induces the sequential unfolding of its three-dimensional structure. Recording the force against tip-surface separation yields a force-distance spectrum characteristic of the unfolding of a single protein. The presence of several distinct events in the force spectrum indicates that secondary structure elements of membrane proteins unfold in well-defined sequences. As their characteristic saw-tooth pattern stems from the extension of already unfolded polypeptide elements, the unfolding spectra are readily analyzed with the wormlike chain (WLC) model. Single-molecule force spectroscopy [14,73,150] provides novel approaches to characterize water-soluble and membrane proteins under variable physiological environments [22,45,46]. In all measurements the proteins were exposed to buffer solution at ambient temperature. It was shown in several examples, that single-molecule force spectroscopy enables to detect inter- and intramolecular interactions within and between proteins [65,79,103,113]. Such experiments not only enabled to detect the stability of membrane proteins [98], but also to probe their energy landscape [60] and refolding kinetics [64]. Single potential barriers confine structurally stable segments that may be represented by transmembrane alpha-helices, polypeptide loops or fragments hereof. These structural segments are established by collective interactions of several amino acids. Once the externally applied force overcomes the stability of these segments they unfold spontaneously. The first experiments allowed investigating how environmental variations such as the oligomeric assembly [115], temperature changes [59], point mutations [98], or pH variations [64] influenced the stability of these structural segments and thereby the unfolding pathways of the protein. Comparing structurally stable segments established within two different membrane proteins having almost identical structures allowed to gain insights into the origin of these interactions [18]. Recently, it has become possible to observe the refolding of secondary structure elements into the final protein and to estimate their folding kinetics from single-molecule experiments [64].

Until now, force-spectroscopy experiments performed on single proteins did not allow to detect and to locate molecular interactions that activate a protein. Moreover, it could not be shown at which location a ligand binds to the protein and thereby activates the protein function. Monitoring the state of activation of a protein would, however, permit to test the effect of test compounds on the functional state of proteins and, thus, allow to more effectively screen for physiologically active drug candidates.

Thus, the technical problem underlying the present invention was to provide methods for determining the functional state of proteins.

The solution to this technical problem is achieved by providing the embodiments characterized in the claims.

Accordingly, the present invention relates to a method for determining the state of activation of a (poly)peptide, comprising the steps of: (a) immobilizing a (poly)peptide on a carrier; (b) attaching the (poly)peptide immobilized on the carrier to a force measuring device of a force spectroscope; (c) applying a pulling force to the (poly)peptide and measuring the force required for stretching and/or unfolding of the (poly)peptide attached to the carrier, wherein the measurements are carried out (i) prior to and (ii) after treatment with a known modulator of said (poly)peptide; (d) comparing the force spectra of the (poly)peptide prior to and after treatment with the modulator; and (e) concluding from a difference of the force spectra of step (d) on the state of activation of the (poly)peptide.

The term the “known modulator” as used herein refers to a compound known to have a particular effect on the state of activation or function of a given (poly)peptide. The known modulator may be for example a known activator, inhibitor, co-factor, substrate or ligand of said (poly)peptide. As explained herein, such a compound—when interacting with the (poly)peptide—will also have an effect on the molecular interactions within the (poly)peptide, stabilizing or destabilizing secondary structure elements (such as alpha-helices or beta-sheets) which will be reflected in the force spectra of the (poly)peptide. In particular cases such a modulator only affects the conformation of the (poly)peptide without having an obvious effect on the activity (of the (poly)peptide (e.g. the conversion rate of an enzyme). In these cases, the modulator may be a cofactor affecting the binding of additional cofactors of the (poly)peptide which in turn may have an effect on the state of activation of the (poly)peptide.

The term “(poly)peptide” refers to peptides, polypeptides and proteins. As used herein, peptides have up to approximately 30 residues, polypeptides between 31 and approximately 50 residues, whereas proteins have at least 51 residues. It is, however, apparent that some small proteins may be made up of a lower number of amino acid residues such as e.g. 35 or 40. The term “(poly)peptide” refers to wild-type (poly)peptide but also to mutants of particular (poly)peptides. It is important to note that the present invention is particularly useful for screening such mutants in order to find out whether or not a particular mutation has an effect on the state of activation of the (poly)peptide. To this end, the force spectra of wild-type and mutant (poly)peptides obtained from measurements in the presence and absence of a known modulator can be compared. If a particular mutation has an effect on e.g. binding of a known activator or if the mutation results in blocking of a conformational change required for activation of the (poly)peptide, this would result in a different force spectrum. The difference would thus indicate to the skilled person that the (poly)peptide mutant no longer is capable of being activated by said known activator.

Immobilizing a (poly)peptide on a carrier can be achieved by covalently coupling the polypeptides to the carrier or by using non-covalent interactions. Non-covalent interactions may e.g. be mediated by antibodies or other biological molecules such as avidin, biotin, histidine tags, collagen binding domains, streptavidin, fibronectin, which are capable of specifically attaching the (poly)peptides, in some cases by interaction with a specific binding partner forming e.g., a biotin/avidin or biotin/streptavidin bridge between the (poly)peptide and the carrier. Other non-covalent interactions resulting in immobilization of the (poly)peptides may be based on hydrophobic, hydrophilic, electrostatic or ionic interactions of the (poly)peptides with the carrier.

The term “force measuring device of a force spectroscope” refers to a cantilever at which the (poly)peptide is attached to. Alternative “force measuring devices” may for example use a dithering tip or a bead or a vesicle to detect molecular forces.

The method of the present invention can be applied to proteins which can exist in at least two states, one of which is an active state or a state which activity has changed in which the (poly)peptide exerts one of its functions. Accordingly, the term “determining the state of activation” means finding out whether a (poly)peptide is in its active state and optionally or alternatively, if so, in which state of activation (see below).

The inactive state is the state in which the protein is e.g. switched off. In case the (poly)peptide is an enzyme, this state may be the state with a reduced or blocked catalytic activity. The inactive state of a (poly)peptide may, however, also be a denatured state of a (poly)peptide. The inactive state may be also presented by the state at which the protein exhibits no functional activity. In case of an ion channel, this may be the state at which the channel is closed.

While some proteins switch between an active and an inactive state, other proteins have multiple states of activation. Allosteric proteins are one example of such proteins which can switch, e.g. upon binding of specific ligands, between a number of states of activation (e.g. states A-→B-→C-→D . . . etc.). Each of these states may have different ligand binding constants. Importantly, the switch between different states of activation is generally accompanied by a conformational rearrangement within the three-dimensional organization of the protein. This structural rearrangement may be limited to only few amino acid residues often located near the active site of the protein, it may affect isolated regions within the secondary structure of the protein (such as an entire alpha helix or beta sheet arrangement) or affect entire subunits of the protein. Such conformational rearrangements, as discussed above, have implications on the internal stability of the proteins, even if only very few amino acid residues are affected. The present invention's methods exploit the idea that each state of activation comes along with a characteristic set of molecular interactions, which on the one hand hold together the three-dimensional organization of the protein and on the other hand drive the protein function. For example, binding of a ligand in the active center of a protein may stabilize the arrangement of the amino acid residues involved in formation of the binding pocket. Accordingly, when such a protein is analyzed by the methods of the present invention, additional pulling force is required for unfolding of the structural elements involved in the formation of the binding pocket. This additional force will be reflected in a change of the force spectrum, in particular in the position of the amino acid residues located near the ligand binding site. Accordingly, the methods of the present invention allow a differentiation between different functional states of proteins. The assignment of a particular state of activation of a (poly)peptide is possible because the methods of the present invention rely on a comparison of the force spectra with a “reference state”. This reference state is the state of the (poly)peptide after treatment with a “reference modulator”, i.e. a modulator with a known effect on the (poly)peptide. A given (poly)peptide is, for example, in its active state when treated under suitable conditions with a known activator. Alternatively, the (poly)peptide is in its inactive state when treated with a compound known to block activation of the (poly)peptide.

Applying the present invention's method, the inventors have been able to generate force curves containing detailed information about strength and location of molecular interactions established within NhaA, an Escherichia coli antiporter specifically involved in exchange of Na⁺ ions for H⁺, allowing the cell to adapt to high environmental salinity and to grow at alkaline pH (Padan et al., 2001). Moreover, these force curves allow to determine the functional state of the NhaA protein in the presence or absence of modulators. As shown in detail in the Examples, while molecular interactions stabilizing secondary structure elements remained unaffected on switching NhaA into its functional (active) state, those being assigned to the Na⁺ binding site changed dramatically. As illustrated by this example, the direct observation of molecular interactions provides novel insights into activation mechanisms of proteins.

In a preferred embodiment of the present invention, the modulator is a known activator, inhibitor, co-factor, substrate or ligand of said (poly)peptide. In fact, this method of the present invention may be performed with any compound with a known effect on the (poly)peptide function. The only constraint, as explained above, is the fact that an assignment to a particular state of activation must be possible.

The present invention also relates to a method for determining the state of activation of a (poly)peptide, comprising the steps of: (a) immobilizing a (poly)peptide on a carrier; (b) attaching the (poly)peptide immobilized on the carrier to a force measuring device of a force spectroscope; (c) applying a pulling force to the (poly)peptide and measuring the force required for stretching and/or unfolding of the (poly)peptide attached to the carrier; (d) comparing the force spectra of the (poly)peptide of (c) with a control representing an inactive or a known state of activation of the (poly)peptide; and (e) concluding from a difference, if any, of the force spectra of step (d) on the state of activation of the (poly)peptide.

The present invention also relates to a method for determining whether a compound is an activator of (poly)peptide function, comprising the steps of: (a) immobilizing a (poly)peptide on a carrier; (b) attaching the (poly)peptide to a force measuring device of a force spectroscope; (c) applying a pulling force to the (poly)peptide and measuring the force required for stretching and/or unfolding of the (poly)peptide attached to the carrier, wherein separate measurements are carried out in the presence of a test and a reference buffer, wherein the test buffer comprises a compound suspected of being an activator of the function of said (poly)peptide and wherein the reference buffer comprises (i) a compound known to have no effect on the activity of said (poly)peptide or (ii) a compound known to be an activator of said (poly)peptide; (d) comparing the force spectra of the (poly)peptide measured in the presence of the test and the reference buffer; and (e) concluding from a difference of the force spectra of step (d) whether the compound suspected of being an activator of said (poly)peptide is an activator of the (poly)peptide.

The term “compound known to have no effect on the activity of said (poly)peptide” refers to a compound which does not affect the function of the (poly)peptide. Such a compound does usually not have an effect on the force spectrum of the (poly)peptide.

The term “activator” refers to a compound having a positive effect on the activity of the (poly)peptide. Hence, such a compound may e.g. increase the catalytic activity of an enzyme, preferably by at least 10% such as at least 30%, more preferably at least 50% such as at least 100%. Most preferred is that the activity is enhanced by at least 200% such as at least 300% or at least 500%. The activator may be e.g. a proteinaceous compound, a peptide or a chemical entity such as a small molecule.

To test for a compound supposed to be an activator of (poly)peptide function, said (poly)peptide is preferably in its inactive state or in an intermediate state out of a plurality of different states of activation where a possible activation can be detected.

Depending on whether reference buffer (i) or (ii) is used in this method, the comparison of the force spectra will yield different results:

When Using Reference Buffer (i):

(a) the observation of no difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer did not interact with the (poly)peptide and/or is no activator of said (poly)peptide.

When Using Reference Buffer (ii):

(a) the observation of no difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer interacted with the (poly)peptide and is an activator of said (poly)peptide.

(b) the observation of a difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer is a modulator, i.e. activator or inhibitor. Said modulator causes the (poly)peptide to (predominantly) adopt a state of activation, possibly out of a plurality of different states of activation as discussed herein above, which differ from the state of activation elicited by said activator comprised in said reference buffer. Depending on the sign of the difference observed, and optionally using prior knowledge regarding the number of distinct states of activation of said (poly)peptide, for example one of the following further conclusions may be drawn: (i) the compound contained in the test buffer interacted with the (poly)peptide and is an inhibitor of said (poly)peptide. This may apply to a case where said (poly)peptide occurs in two states, namely an active and an inactive state. (ii) the compound contained in the test buffer interacted with the (poly)peptide and is an activator of said (poly)peptide. This may apply to a case where said (poly)peptide occurs in at least three states and the state of activation elicited by the compound contained in the reference buffer is an intermediate state of activation different from the one elicited by the compound contained in the test buffer

The above described screening assay represents an adaptation of the first assay described in the present invention and allows to screen compounds with respect to their capability to modulate the state of activation of a given (poly)peptide. According to this method, the state of activation of a (poly)peptide is monitored in the presence and absence of a potential activator of said (poly)peptide. Binding of an activating compound will induce a change of the force spectra which is indicative of the compound's capability to modulate the function of the (poly)peptide.

It is important to note that some proteins can bind specific activators as well as other specific cofactors required for optimal protein function. In fact, the binding constant of an activator of a protein may be affected from binding of a cofactor. In such cases, potential activators may also be screened after treatment and/or in the presence of a known specific cofactor.

The present invention also relates to a method for determining whether a compound is an inhibitor of (poly)peptide function, comprising the steps of: (a) immobilizing a (poly)peptide on a carrier; (b) attaching the (poly)peptide to a force measuring device of a force spectroscope; (c) applying a pulling force to the (poly)peptide and measuring the force required for stretching and/or unfolding of the (poly)peptide attached to the carrier, wherein separate measurements are carried out in the presence of a test and a reference buffer, wherein the test buffer comprises a compound suspected of being an inhibitor of the function of said (poly)peptide and wherein the reference buffer comprises (i) a compound known to have no effect on the activity of said (poly)peptide; (ii) a compound known to be an activator of said (poly)peptide; (iii) or a compound known to be an inhibitor of said (poly)peptide; (d) comparing the force spectra of the (poly)peptide measured in the presence of the test and the reference buffer; and (e) concluding from a difference of the force spectra of step (d) whether the compound suspected of being an inhibitor of said (poly)peptide is an inhibitor of the (poly)peptide.

The term “inhibitor” refers to a compound having a negative effect on the activity of the (poly)peptide. Hence, such a compound may e.g. reduce or block the catalytic activity of an enzyme, preferably by at least 10% such as at least 30%, more preferably at least 50% such as at least 100%. The inhibitor may be e.g. a proteinaceous compound, a peptide or a chemical entity such as a small molecule, it may be isolated from nature or be a synthetic compound. To test for a compound supposed to be an inhibitor of (poly)peptide function, said (poly)peptide is preferably in its active state or in an intermediate state out of a plurality of different states of activation where a possible inhibition can be detected.

When Using Reference Buffer (i):

(b) the observation of a difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer interacted with the (poly)peptide and is an inhibitor or activator of said (poly)peptide depending on the initial state of activation of said (poly)peptide.

When Using Reference Buffer (ii):

(a) the observation of no difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer interacted with the (poly)peptide and is an activator of said (poly)peptide

(b) the observation of a difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer is a modulator, i.e. activator or inhibitor. Said modulator causes the (poly)peptide to (predominantly) adopt a state of activation, possibly out of a plurality of different states of activation, which differ from the state of activation elicited by said activator comprised in said reference buffer. Depending on the sign of the difference and optionally using prior knowledge regarding the number of distinct states of activation of said (poly)peptide, for example one of the following further conclusions may be drawn: (i) the compound contained in the test buffer interacted with the (poly)peptide and is an inhibitor of said (poly)peptide. This may apply to a case where said (poly)peptide occurs in two states, namely an active and an inactive state. (ii) the compound contained in the test buffer interacted with the (poly)peptide and is an activator of said (poly)peptide. This may apply to a case where said (poly)peptide occurs in at least three states and the state of activation elicited by the compound contained in the reference buffer is an intermediate state of activation different from the one elicited by the compound contained in the test buffer.

When Using Reference Buffer (iii):

(a) the observation of no difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer interacted with the (poly)peptide and is an inhibitor of said (poly)peptide

(b) the observation of a difference between the force spectra generated by the test and the reference buffer of step (d) will allow to conclude that the compound contained in the test buffer is a modulator, i.e. activator or inhibitor. Said modulator causes the (poly)peptide to (predominantly) adopt a state of activation, possibly out of a plurality of different states of activation, which differ from the state of activation elicited by said inhibitor comprised in said reference buffer. Depending on the sign of the difference and optionally using prior knowledge regarding the number of distinct states of activation of said (poly)peptide, for example one of the following further conclusions may be drawn: (i) the compound contained in the test buffer did not interact with the (poly)peptide. This may apply to a case where said (poly)peptide occurs in two states, namely an active and an inactive state. (ii) the compound contained in the test buffer interacted with the (poly)peptide and is an activator of said (poly)peptide. This may apply to a case where said (poly)peptide occurs in two states, namely an active and an inactive state. (iii) the compound contained in the test buffer interacted with the (poly)peptide and is an activator of said (poly)peptide. This may apply to a case where said (poly)peptide occurs in at least three states and the state of activation in which said (poly)peptide occurred at the beginning of the measurement was an intermediate one (iv) the compound contained in the test buffer did not interact with the (poly)peptide. This may apply to a case where said (poly)peptide occurs in at least three states and the state of activation in which said (poly)peptide occurred at the beginning of the measurement was an intermediate one.

As shown in the Examples, from comparing the force spectra of the test and the reference buffer it is also possible to determine the amino acid residues of the (poly)peptide that interacted with the compound contained in the reference buffer. The characteristic force spectroscopy spectra revealed unfolding NhaA contains sets of intensive force peaks. To assign the molecular interactions that were established within the protein each force peak of the force spectra is fitted using the worm-like chain (WLC) model. The fit revealed the length of the stretched polypeptide and allowed to assign polypeptide regions which formed molecular interactions. If a compound interacts with a certain region of the (poly)peptide this will change molecular interactions which are then measured by the force spectroscopy experiments. The additional or changed force peak caused due to the compound interaction will be located and quantified using the above procedure.

The above described screening assay represents an adaptation of the first assay described in the present invention and allows to screen compounds with respect to their capability to inhibit the activity of a given (poly)peptide. According to this method, the state of activation of the (poly)peptide is monitored in the presence and absence of a potential inhibitor of said (poly)peptide. Binding of an inhibiting compound will induce a change of the force spectra which is indicative of the compound's capability to modulate the function of the (poly)peptide.

It is important to note that some proteins can bind specific activators as well as specific inhibitors of their protein function. In fact, some proteins can bind a number of additional factors, all of which may affect substrate binding and/or binding of the individual factors. Hence, the binding constant of a second binding partner of the protein (e.g. an inhibitor) may be affected from binding of a first modulator (e.g. an activator). In such cases, potential inhibitors may also be screened after treatment and/or in the presence of other known binding partners of said protein, such as specific activators or specific cofactors of the protein.

In a preferred embodiment of the present invention the force-distance spectra recorded when applying any one of the above mentioned reference buffers to the (poly)peptide are deposited in any suitable format, preferably in a database. This applies to all of the following embodiments.

In a preferred embodiment of the present invention, the carrier is (a) a biological carrier selected from the group consisting of membrane, vesicle, cell, interface such as cell membranes, membrane of vesicles, lipid membranes, extracellular matrix; biological structure such as lipids, collagen, actin, microtubules, cytoskeleton, protein and protein complexes, aggregates such as amyloid fibers and plaques, bone, nucleic acid molecule, fibril, fiber, and biological scaffold (tissues, cells) or (b) a non-biological carrier selected from the group consisting of solid state material such as mica, graphite, silicium, gold, gallium arsenide; polymer such as polyethylene, polypropylene, polystyrol, acrylate, collagen; synthetic membrane such as co-block polymers, or membrane assembled from synthetic lipids; and synthetic scaffold such as peptide hydrogels, hydrogels, polymer nanofibers, peg-pamam star polymers.

In another preferred embodiment of the present invention, the (poly)peptide is arranged as a three-dimensional crystal, two-dimensional crystal, bound to a biological or non-biological surface (e.g. solid state material, polymer, synthetic membrane, synthetic scaffold), attached to a membrane or peptide, attached to any biological or synthetic molecule (e.g. amines, antibiotics, hormones, enigmols, supramolecular cylinders of formula [Fe₂L₃]⁴⁺), or incorporated into a membrane. The (poly)peptide may be attached or bound covalently (e.g. using C- or S-bonds) or non-covalently (e.g. via electrostatic, hydrophilic, hydrophobic, ionic, electrostatic, magnetic or van der Waals interactions).

In another preferred embodiment of the present invention, the (poly)peptide is embedded in a lipid bilayer. The lipid bilayer may be derived (i.e. obtainable) from a biological membrane. Alternatively, the lipid bilayer may be a synthetic lipid bilayer. “Synthetic” means that it is generated from its constituents in vitro. To this end, molecules such as phospholipids, sphingolipids, glycolipids, cholosterol and other biological or synthetic molecules may be mixed in the presence or after addition of the (poly)peptide.

In yet another preferred embodiment of the present invention, the measuring device is selected from (a) tip of a force spectroscope, (b) a magnetic tweezer, (c) an optical tweezer, (d) a protein or protein complex, (e) any other biological or synthetic molecule such as a peptide, protein, protein complex, lipid, nucleic acid, (D surface of a force apparatus such as mica, graphite, silicium, gold, gallium arsenide, (g) a membrane such as a cell membrane, vesicle, lipid bilayer, lipid monolayer, co-block polymer, surface layer of a bacterium or vertebrate cell, and (h) the probe of a scanning probe microscope or a scanning probe spectroscope.

In another preferred embodiment of the present invention, the (poly)peptide is attached to the measuring device by non-covalent interactions. The term “non-covalent interaction”, as used throughout the specification means hydrophobic, hydrophilic or steric interactions or/and interactions by charge (e.g. van der Waals, ionic, electrostatic). To this end, the measuring device and the (poly)peptide are brought into contact. Applying a force of 300 to 1000 pN results in binding of one terminal end to the measuring device, preferably either via a covalent bond or enforced non-specific adsorption.

In another preferred embodiment of the present invention, the pulling force applied from the tip of the force spectroscope is in the range of 1 to 500 pN. The maximum pulling force is determined by the strength of the molecular interaction established within the protein. The molecular interaction can also result from the binding of the compound.

In another preferred embodiment of the present invention, stretching and/or unfolding are measured by recording the unfolding and/or stretching resistance observed upon application of the pulling force from the tip of the force spectroscope to the (poly)peptide. It is to be noted that the force applied to a single protein plays the role of a denaturant leading from continuous extension of a protein to complete unfolding of its three-dimensional structure. The force required for withdrawing the measuring device attached to the (poly)peptide from the carrier represents the pulling force. As soon as this pulling force exceeds the stability of the protein, it induces the sequential unfolding of its three-dimensional structure which is recorded.

In another preferred embodiment of the present invention, said activator is an agonist. The term “agonist” refers to analogues of a native or synthetic ligand (for example a protein or a hormone) that binds to a specific receptor and triggers the receptor activity function. It does not matter whether the ligand is a natural or synthetic compound. Most important however, is that the binding of the ligand induces a biological reaction, such as the activation of a receptor, which then can activate a biological process. One example is given by the acetylcholine receptor, which opens the channel after binding of the agonist acetylcholine. The term “antagonist” describes analogues that act as competitive inhibitors against agonist binding. They may also displace the agonist from the receptor and occupy the appropriate receptor. As a result the antagonist prevents receptor activation or changes the receptor function.

In another preferred embodiment of the present invention, said activator or inhibitor is selected from the group consisting of a protein specific ligand (for example a protein or a hormone), a synthetic compound such as a synthetic ligand, ligand, activator, agonist, antagonist; of acetylcholine, nicotine, glutamate, dopamine, hydroxytryptamine, serotonin, pheromones, interleukin and synthetic analogues or substitutions thereof; a pharmaceutical compound such as a hormone or a toxin (e.g. one of the above mentioned compounds); a biochemical or biological compound such as mentioned above or a, (poly)peptide, cholesterol, lipid, signaling molecule, light, electrolyte, pH, voltage or current, being capable of inducing a change of the functional state (activity) of said (poly)peptide.

In an additional preferred embodiment of the present invention, more than one compound suspected of being an activator or inhibitor of the (poly)peptide function is tested.

In another preferred embodiment of the present invention, the measurement is performed in the presence of a compound which is an agonist or antagonist of said activator or inhibitor in order to show if said activator or inhibitor of (poly)peptide function is modulated by said agonist or antagonist.

In a more preferred embodiment of the present invention, said compound is part of a compound library and said method contains the additional step of screening said compound library for identifying a lead compound for drug development.

In another more preferred embodiment of the present invention, said screening is high-throughput screening carried out by multiple cantilevers and/or by multiple force spectroscopes and/or by fast-speed force spectroscopy.

In a preferred embodiment of the present invention, the (poly)peptide is a protein.

In a more preferred embodiment of the present invention, said protein is selected from the group consisting of (a) ligand-gated receptor, (b) G-protein coupled receptor, (c) ion channel, (d) water channel, (e) antiporter, (f) communication channel, (g) symporter, (h) porin, (i) ion gated channel, electrolyte gated channel and pore, ion pump, glutamate gated ion channel, ATP gated channel, (j) mechanosensitive channel and (k) ATP synthase.

Ligand-gated receptors include for example nicotinic acetylcholine receptor, hydroxytryptamine serotonin receptor, GABA A receptor, GABA B receptor. G-protein coupled receptors include molecules such as rhodopsin, vasopressin V2 receptor, metabotropic glutamate receptors, interleukin-8 receptor, adrenergic receptors, neuropeptide Y receptor, dopamine 1B receptor, glycoprotein hormone receptor, melanocortin receptor, adenosine receptor, pheromone A receptor. Ion channels include molecules such as potassium channel, sodium channel, calcium channel, slow voltage-gated potassium channel, NMDA receptor, P2X5 purinoceptor, chloride channel CLC, influenza virus matrix protein M2, calsequestrin, arsenical pump membrane protein, mechanosensitive ion channels MscS and MscL, voltage-dependent calcium channel. Water channels include molecules such as aquaporin 1-10, MIP, GIpF. Pores include for example channel forming colicins, ATP P2X receptor, delta-endotoxin CytB, P2X purinoceptors, mammalian defensin, proteinase inhibitor 117, Hemolysin E. Antiporters include for example sodium/proton antiporter, betaine/proton antiporter, cadmium-transporting ATPase, anion exchange protein, sodium/calcium exchanger, calcium/proton exchanger, sodium/hydrogen exchanger, multicomponent K⁺:H⁺ antiporter, K⁺-dependent Na⁺/Ca⁺ exchanger, multicomponent Na⁺:H⁺ antiporter, sodium/hydrogen exchanger. Communication channels include for example gap junctions, connexins. Symporters include for example LacY, sodium:alanine symporter, Na⁺ dependent nucleoside transporter, glycine neurotransmitter transporter, sodium/glutamate symporter, pentulose kinase. Porins include for example OmpF porin, maltoporin. Additional (poly)peptides or proteins that may be analyzed using the methods of the present invention are mentioned above (points i and k) including ATP synthases of e.g. F-, V-, or H-type.

In a preferred embodiment of the present invention, the pulling force is exerted by the measuring device. In this case the cantilever (if measuring device) is moved to stretch the protein.

In another preferred embodiment of the present invention, the pulling force is exerted by the carrier. In this case the carrier is moved to stretch the protein.

In another preferred embodiment of the present invention, the pulling force is exerted by the (poly)peptide. In this case the (poly)peptide exerts forces that deflect the cantilever. Positions of the cantilever and carrier are not changed in this case.

In another preferred embodiment of the present invention the pulling force is exerted by applying two or three of the above embodiments.

The figures show:

FIG. 1:

Detecting pH and Na⁺ dependence of molecular interactions established within NhaA. (A) Left, representative force-extension curve recorded upon mechanical unfolding a single NhaA molecule. Each force peak is fitted by the WLC model (solid curves) with the numbers of stretched aa residues given. Right, secondary structure of NhaA mapped with stable structural segments detected (grey shaded) upon pulling the C-terminus. Grey gradients reflect uncertainties in determining segment positions. To determine merges of the segments on the opposite side to the AFM tip of the membrane or within, the membrane thickness of 4 nm was considered (Kedrov, 2004). In these cases corresponding contour lengths are given in brackets. (B-F), superimpositions of extension curves recorded upon single NhaA unfolding at pH 3.8 (B) and (C) 5.5 (inactive states) and (D) pH 7.7 (active state) at electrolyte concentrations of 150 mM KCl and 50 mM NaCl. The pH-dependent unfolding peak at 225aa is encircled. To prove the reversibility of the pH dependent change NhaA was incubated for 1 h at pH 7.7 and unfolded at pH 3.8 (E). Significant restoration of the protein stability suggests the reversibility of molecular interactions. (F) Force-extension curves of NhaA recorded at pH 7.7 in absence of NaCl reduced the molecular interaction to that measured for the inactive state. 20 force-extension curves were superimposed for each figure. (G) Average unfolding forces of helical pairs. Forces and standard deviations are plotted for different pH values at 150 mM KCl and 50 mM NaCl. Pairs of neighboring helices tend to unfold cooperatively giving force peaks at 163aa (helices VII&VIII), 202aa (helices V&VI), 258aa (helices III&IV) and 328aa (helices I&II)(Kedrov, 2004).

FIG. 2:

pH and Na⁺ dependent molecular interactions established at the active site of NhaA. Change in stability of helix V derived from single-molecule unfolding events. Distribution of unfolding forces of helix V (peak 225aa) in presence of Na⁺ is given by histograms for pH 3.8 (A), 5.5 (B), and 7.7 (C). (D) Removal of Na⁺ ions from the buffer solution reduced the molecular interaction to that measured for inactive NhaA (A and B). Approximately 70 single-molecule unfolding spectra were analyzed at each pH (for methodological details see Example 5). Distributions A and C, C and D are statistically different with significance of p<0.001. Black bars on the left of the histograms represent unfolding events, where no peak was detected at 225aa.

FIG. 3:

Characterizing molecular forces established at ligand binding site of NhaA. Strength (A) and frequency ¹(B) of molecular interactions established at ligand binding site (C) increase upon changing pH from 5 to 6. Solid lines represent sigmoid fits of the data points. Dashed lines indicate pH values at which the mid-points of transitions were reached. (C) Primary and secondary structure of helix V. Aspartic acids (D163 and 164) of the Na⁺ binding site were indicated. pH changes enable accessibility of Na⁺ ions. ¹the term “frequency” as used herein means probability of detecting a peak

FIG. 4:

Inhibitor binding changes molecular interactions within NhaA. Superimpositions of 20 NhaA force-distance spectra recorded in absence (a) and in presence (b) of 2-aminoperimidine (AP), 190 mM KCl, 10 mM NaCl, pH 7.7. The local change of molecular interactions (encircled) was located at α-helix IX. Distribution of forces detected at α-helix IX at 0 (c), 20 (d), and 40 μM (e) AP. Two populations of NhaA molecules were detected in presence of AP. Weak interactions (left Gauss fit) were observed for free NhaA and enhanced interactions (right Gauss fit) for the NhaA.AP complex. ¹the term “frequency” as used herein means probability of detecting a peak

FIG. 5:

Histograms of the α-helix IX stability reflect a reduced AP binding due to excess of a competing substrate (a) and pH-locked conformation (b) of NhaA.

The examples illustrate the invention:

EXAMPLE 1
NhaA Remains Fully Folded Over Wide pH Range

FIGS. 1B-D show superimpositions of force-extension curves recorded upon unfolding individual NhaA molecules at different pH values. They correspond to unfolding of inactive (pH 3.8 and 5.5) and fully active (pH 7.7) forms of NhaA (Taglicht et al., 1991). All superimpositions show the characteristic unfolding spectra of NhaA as reported (Kedrov, 2004) and reveal no additional unfolding events. The unfolding peaks detected allow locating structural segments, which formed an unfolding barrier (FIG. 1A). Overcoming the critical force initiates cooperative unfolding of all aa within the structural segment, which established the stabilizing molecular interactions (Kedrov, 2004). These stable segments do not necessarily correlate to a single secondary structure element of the protein. For example, the force spectra showed unfolding of a transmembrane helix together with a polypeptide loop, or of two helices collectively establishing an unfolding barrier (FIG. 1A). Each barrier then unfolds cooperatively upon mechanical pulling.

Average forces required to unfold helical pairs I&II, III&IV, V&VI and VII&VIII (FIG. 1G) show that their stability is retained independent of the pH range of 3.8 to 7.7. Both, the unaffected stability and locations of molecular interactions stabilizing the structural domains of NhaA imply that they do not change upon protein activation. Hence, it can be concluded that the protein maintained its folded stable conformation in the experiments.

EXAMPLE 2
Alpha-Helix V Establishes Molecular Interactions

While the general profile of NhaA unfolding curves remained unchanged upon pH variation, the molecular interactions establishing the force peak 225aa increased significantly (FIG. 1B-D; encircled areas). On the basis of the primary and presumed secondary structure of NhaA (Rothman et al., 1996), we recently showed that the corresponding unfolding barrier was located at the middle of transmembrane helix V (FIG. 1A) (Kedrov, 2004). Overcoming the molecular interactions stabilizing this structural region by an externally applied force induces unfolding of the cytoplasmic half of helix V. On raising the pH from 3.8 to 7.7, the average force required to overcome the molecular interactions increased from 74±29 pN to 107±26 pN (average ±SD). Thus, in fully active NhaA the molecular interactions established within this region reached the strength typically measured for unfolding of a helical pair (FIG. 1G, 3A). Simultaneously, the frequency of peak detection increased from 31 to 94% (FIG. 3B). Detailed insights into the kinetics of this local stabilization were achieved analyzing single-molecule unfolding events (FIG. 2A-C). The histograms of the unfolding forces distribution clearly show an increased frequency of the 225aa peak while approaching functional pH values. The increased stability of this region shifted the force distribution gradually to ˜100-120 pN. Reversing the pH from 7.7 to 3.8 restored initial molecular interactions (FIG. 1E) as the mean unfolding force reduced to values (78±30 pN) similar to that detected for the inactive form of NhaA (74±29 pN).

EXAMPLE 3
pH Dependent Molecular Interactions Co-Localize with Ligand-Binding Site

Several studies on NhaA imply that the negatively charged aspartic acid residues 163 and 164 are involved in the Na⁺-binding site located in the center of transmembrane helix V. Substitution of these residues with cysteines or asparagines dramatically reduce the cation transport activity of NhaA (Inoue et al., 1995; Padan et al., 2001). It has been shown for different membrane proteins that ligand binding during the functional cycle can alter the protein conformation (Ferguson et al., 2002; le Coutre et al., 2002; Wang, 1997) causing changes in molecular interactions. As the observed change in interactions is localized in the direct proximity of the ligand-binding site of NhaA, we applied force spectroscopy to probe the effect of Na⁺ ions on this site. For this purpose we unfolded single NhaA molecules at pH 7.7 in absence of NaCl (FIG. 1F). To eliminate possible effects of non-specific electrostatic interactions, the total ionic strength of the buffer solution was kept constant using KCl as substitute. The force experiments did not detect any change of the unfolding pathways upon absence of Na⁺ ions except for the 225aa peak, which almost disappeared. Moreover, the distribution of unfolding forces (FIG. 2D) exhibited two peaks at 60-70 pN and 90-100 pN. The peak at 90-100 pN suggested that a certain fraction of the molecules (30-35%) still possessed strong molecular interactions within helix V. The specificity of Na⁺ ions strongly suggests that they play a major role in establishing the molecular interactions at the ligand binding site of helix V. Together with the observed pH-dependent formation of these interactions it may be suggested that the accessibility of the binding site to Na⁺ ions is governed by pH. As shown recently, this ion binding capability can be altered upon small changes of the side-chain orientation (Wang, 1997), which is promoted by minute spatial rearrangements of NhaA helices. Thus we conclude that the antiporter is activated by intramolecular interactions, which are established only at neutral pH and simultaneously occurring ligand binding.

EXAMPLE 4
Molecular Interactions Establish Full Strength at High Probability to Activate Ion Channel

To analyze formation and kinetics of molecular interactions that were established upon NhaA activation the frequency of peak appearance at aa 225 and rupture forces were measured at pH values ranging from 3.8 to 7.7. The data suggested that the interactions gradually increased when the pH was increased from 5 to 6 (FIG. 3A). The probability of the peak appearance continuously increased with the pH as well (FIG. 3B). It showed, however, a much lower slope beginning at pH 4 (˜30%) and finally reaching ˜95% at pH 7.5. Sigmoid distributions (FIGS. 3A, B; black curves) accurately fitted the data points assuming C1=39, C2=70 for the force and C1=29, C2=63 for the probability. Mid-points of both transitions were located at pH0=5.4 (force) and 5.7 (probability). Clearly, both mid-points for establishing the molecular interactions were shifted to the acidic range, compared to pH 7.5 optimum reported for NhaA activity (Taglicht et al., 1991). One possible explanation could be that the observed formation of molecular interactions within transmembrane helix V relates to an early activation step of the protein, while previously reported structural changes (Rothman et al., 1997; Venturi et al., 2000) finalize the activation of the antiporter at pH range 7-8. The full activity of NhaA would then be reached at pH 7.0 at which the molecular interactions at the active site of the protein reached their full strength and occurred with a probability of >90%. Thus, we conclude that establishing the full strength of molecular interactions builds an initial step towards activating a single NhaA. These observations are in agreement with those made on lactose permease (Abramson, 2003; Sun, 1998; Zhang, 2002), a paradigmatic secondary transporter (Abramson, 2004). Here it was shown that activation of this transporter is a multi-stage process. Thus, the observed molecular interactions may represent initial steps of conformational changes for NhaA, which have been previously observed at pH ranging between 7 and 8 (Rothman et al., 1997; Venturi et al., 2000). To activate all NhaA channels these molecular interactions have to be established in every NhaA molecule. This finding revealed by single molecule studies complements conventional experiments revealed from large protein assemblies. We conclude that apparently some proteins were activated by establishing their full strength of molecular interactions while other were not activated.

The unfolding spectra of membrane proteins significantly depend on the loading rate which reflects the non-equilibrium nature of the single-molecule experiments. As a result, the frequency of side peaks appearance can be modulated by pulling speed (Janovjak, 2004). Thus, we performed additional unfolding experiments on inactive NhaA (pH 3.8) at higher pulling speed of 1 μm/s, which shifted the frequency of the 225aa peak to ˜55%. In contrast, at pH 7.7 the frequency remained ˜95% (data available upon request). Remarkably, irrespective of the pulling speed the molecular interaction centered at aa 225 reached its maximum stability at pH ˜6. Thus, we assume that changing the pulling speed shifts the slope of the molecular forces and of their probability to occur within the active site of NhaA. The conclusions remained unchanged, however, that the full strength of interactions is established at a pH range before NhaA becomes fully active.

EXAMPLE 5
Inhibitor Binding Establishes Interactions Different to Those Induced by Ligand Binding

We applied the specific NhaA inhibitor 2-aminoperimidine (AP) to NhaA activated at pH 7.7. The force-distance recorded spectra with a SMFS (single-molecule force spectroscopy) exhibited all characteristic peaks from active NhaA (FIGS. 4 A and B). The only significant change was observed for the interaction established at helix XI, which was detected by the 125aa force peak (FIGS. 4 A and B encircled areas). B addition of the inhibitor the average strength of this interaction increased from 75±29 pN (mean ±SD, n=61) to 105±41 pN (n=95), p<0.0001 in presence of 40 μM AP. Histograms showing the distribution of the unfolding forces of the 125aa peak reveal further insights into the localized molecular interactions changing after AP binding. In the absence of AP most NhaA molecules (>90%) established local interactions of ≈75pN (FIG. 4 C). After exposure to AP, a fraction of NhaA molecules enhanced their interaction strength to ≈140 pN (FIG. 4 C,D). This observation suggests that the inhibitor binding established enhanced interactions within the NhaA.AP complex, which were different to those induced by the ligand binding to active NhaA. When assuming a competitive inhibition mechanism the AP binding should be reduced at enhanced Na⁺ concentration. To test this hypothesis we studied the NhaA.AP interactions in presence of 10 or 200 μM NaCl at 40 μM AP. Histograms of the force established at helix IX (FIG. 5 A) showed a drastic decrease of the “stable” fraction, i.e. the AP binding was suppressed with increasing Na⁺ concentration. To study inhibitor interactions with the pH-locked conformation (Taglicht et al., 1991) NhaA was exposed to pH 4.0, which ensures closure of the ion-binding pocket, and then incubated with 40 μM AP. The experiments detected much less NhaA molecules ≈15% that have established enhanced an enhanced stability at the α-helix IX (FIG. 5 B). Thus we conclude, that the protein-inhibitor interactions were hindered in the locked conformation of NhaA.

EXAMPLE 6

In the following, the methods for carrying out examples 1 to 4 are described:

NhaA Preparation

NhaA was overexpressed in E. coli BL21(DE3) with a His₆-tag fused to the C-terminus (Olami et al., 1997). Purification and 2D crystallization was performed at pH 4 (Williams et al., 1999), where the molecule is inactive. The tubular 2D crystals exhibited unit cell dimensions of 48 Å×181 Å with a p22₁2₁symmetry. NhaA activity was determined measuring the active transport of Na⁺ ions using the electrophysiological method of solid supported membranes (Seifert, 1993). The measured pH-profile of activation was identical to that obtained by transport measurements of ²²Na (Taglicht et al., 1991).

AFM

The AFM used (Nanoscope IIIa) was equipped with a fluid cell and 200 μm long Si₃N₄AFM cantilevers (di-Veeco, Santa Barbara). Spring constants of cantilevers were determined (≈0.06N/m) using the equipartition theorem (Butt H. J., 1995; Florin, 1995). 2D crystals of NhaA were immobilized on freshly cleaved mica in 150 mM KCl, 10% glycerol, 25 mM K⁺-acetate, pH4 for 20 min. Experiments were performed in buffer solutions containing 150 mM KCl and 50 mM NaCl at pH 3.8 (20 mM citric acid), 4.5 (20 mM K⁺-acetate), 5.0 (20 mM citric acid), 5.5 (20 mM MES), 6.3 (20 mM HEPES), 7.1 (20 mM HEPES), and 7.7 (20 mM Tris). Na⁺-free buffers contained less than 0.1 mM Na⁺ as estimated by atomic absorption spectroscopy. All buffer solutions were made in fresh nano-pure water (18.2 MΩ·cm), using reagents from Sigma/Merck of p.a. purity grade. Upon buffer exchange the setup was equilibrated for 30 min. After AFM imaging immobilized crystal patches (Kedrov, 2004) an unperturbed area was selected to unfold individual proteins. The AFM tip was then brought in contact with the protein applying a force of 0.5-1 nN to attach its terminal end. After 1 s the tip was withdrawn from the membrane at 120 nm/s, while the cantilever deflection was detected. The value of the deflection at each time point was used to calculate the force acting on the molecule via Hook's law.

Data Analysis

Pulling NhaA (402aa) from either the N- or C-terminus yielded characteristic force-extension curves each exhibiting a length of ˜100 nm (Kedrov, 2004). In this study we focused on C-terminal unfolding events because of dramatic decrease in frequency of N-terminal unfolding events observed at higher pH. Force-extension curves recorded upon single-protein unfolding were manually superimposed. To obtain the unfolded polypeptide chain length each peak was fitted using the worm-like-chain (WLC) model (Bustamante C., 1994) as described (Kedrov, 2004).

Percent probability and average unfolding forces were calculated for each force peak. The standard error of the mean frequency value was derived from the binomial distribution. We analyzed 74 (pH 3.8), 43 (pH 4.5), 60 (pH 5.0), 70 (pH 5.5), 59 (pH 6.2), 68 (pH 7.1), 74 (pH 7.7) events at the pH indicated. Distributions of unfolding force and percent probabilities vs. pH were fitted using the sigmoid function described by

$f (pH) = C_{1} + \frac{C_{2}}{1 + \log^{- (pH - {pH}_{0})}}$

C₁, C₂determine the limits of the function at low and high pH values, pH₀the mid-point of the transition.

EXAMPLE 7

In the following the methods for performing example 5 as different from the methods for performing examples 14 are described:

SMFS

We used a NanoWizard AFM (JPK Instruments, Germany) with an 850 nm laser detection system. As AP absorbs light between 290 and 350 nm, we avoided double-photon excitation and photobleaching by the laser. The spring constants of the 200 μm long (di-Veeco, USA) and 80μ long (Olympus, Japan) Si₃N₄AFM cantilevers used were determined using the equipartion theorem (Butt H. J., 1995; Florin, 1995). NhaA was reconstituted into E. coli polar lipid bilayers forming two-dimensional crystals (Williams et al., 1999). Membranes were immobilized on freshly cleaved mica in 200 mM KCl, 10% glycerol, 25 mM K⁺-acetate, pH 4.0 for 20 min. Experiments were performed in buffer solutions containing 190 mM KCl and 10 mM NaCl at pH 7.7 (20 mM tris-HCl) and pH 4.0 (20 mM citric acid). AP concentrations of prepared solutions were determined by their optical densities at λ=305.5 using the excitation coefficient eAP=7500 M⁻¹·cm⁻¹(Additional Ref. 151). Solutions containing AP were kept away from light and experiments were performed in a dark room. Buffer solutions were made in fresh nano-pure water (18.2MΩ·cm⁻¹), using p. a. purity grade reagents from Sigma/Merck. AFM topographs of immobilized membrane patches resolved the crystalline NhaA arrangements, which were selected to unfold individual proteins (Kedrov et al., 2004).

Data Analysis

Pulling NhaA (402aa) from either the N- or C-terminus yielded characteristic force-distance curves each exhibiting a length of ≈100 nm. Only C-terminal unfolding events were studied as they are dominant at pH 7.7 (Additional Ref. 152). To obtain the unfolded polpeptide chain length each force peak was fifted using the worm-like-chain (Bustamante et al., 1994) model as described (Kedrov et al., 1994). Individual unfolding barriers were located by subtracting the unfolded polypeptide length fron the C-terminus. For each force peak (unfolding barrier) the probability and average forces were calculated. At every studied condition up to 150 single-molecule unfolding events were analyzed using Igor Pro (Wavemetrics Inc., USA) and home-written macros. To probe the statistical difference of data sets the data were tested against one-way ANOVA analysis.

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Method of Determining the State of Activation of a Protein

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