From the Guelph-Waterloo Centre for Graduate Work in Chemistry and Biochemistry, Department of Chemistry and Biochemistry, University of Guelph, Guelph, Ontario N1G 2W1, Canada
Received for publication, March 7, 2003 , and in revised form, April 15, 2003.
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Colicin E1 is a member of the channel-forming subfamily of colicins and is secreted by E. coli that possesses the naturally occurring colE1 plasmid; the holoprotein consists of three functional segments, the translocation, receptor-binding, and channel-forming domains. Initially, the receptor-binding domain (8) interacts with the vitamin B12 receptor of target cells. Following recognition, the translocation domain associates with the tolA gene product, which permits the translocation of colicin E1 across the outer membrane and into the periplasm (9). In the periplasm, the channel domain undergoes a conformational change to an insertion-competent state and then inserts spontaneously into the cytoplasmic membrane of the host cell, forming an ion channel. The channel allows the passage of monovalent ions, resulting in the dissipation of the cationic gradients (H+, K+, and Na+) of the target cell, causing depolarization of the cytoplasmic membrane. In an effort to compensate for the membrane depolarization effected by the colicin E1 channel, Na+/K+-ATPase activity is increased in the host cell, resulting in the consumption of ATP reserves, without concomitant replenishment (4). The final outcome is host cell death.
In addition to structural similarities between colicin E1 and several membrane-targeting proteins, an intriguing characteristic shared by colicin E1 and many membrane-active proteins is the observed requirement of an acidic environment for activation of the soluble structure in order to bind and insert into target cell membranes. The important effect of an acidic environment in conferring an optimum activity in vitro, and perhaps in vivo, for colicin E1 has been attributed to an onset of acid-induced protein unfolding events, resulting in an increased structural mobility/flexibility that may potentiate the complex sequence of structural changes of the channel peptide observed at the surface of a membrane (1012).
An important contribution to our understanding of the mechanism of colicin E1 low pH activation was conducted by Merrill et al. (13) in which a precise pH-sensitive region within the soluble channel domain was identified. By using solution steady-state and time-resolved fluorescence measurements of single Trp mutant channel peptides, Merrill et al. (13) demonstrated that only two of the nine Trp residues engineered at different sites throughout the protein, Trp-413 and Trp-424, exhibited pH-sensitive fluorescence parameters. The fluorescence quantum yield and the lifetime decay components of Trp-413 and Trp-424 exhibited significant pH-sensitive changes that indicated an altered Trp chemical environment as a consequence of site-specific structural changes induced within the channel domain upon acidification.
These observations led to the proposal that the peptide segment encompassing residues 413 and 424 (helices 4 and 5a) within the colicin E1 channel domain undergoes a helix-to-coil transition during solution acidification. This low pH-induced unraveling of helices 4 and 5a was proposed to result from the disruption of a critical salt bridge and three hydrogen bonds that stabilize the secondary structure of this segment of the protein (6, 13). The disruption of one or all of these H-bond links may provide the initial driving force for a cascade of structural events within the channel peptide that lead to the activated, membrane-competent state of the protein.
In the present work, we report the preparation of single and double Asp-to-Ser mutations localized to helices 4 and 5a of the colicin E1 channel domain, which is believed to be the heart of the pH trigger motif (13). Our findings show that Asp-to-Ser mutations disrupt critical H-bonds within the trigger motif and induce a helix-to-coil transition. Furthermore, we demonstrate the effect of these mutations on the activation mechanism (function) of the channel domain by measuring channel domain binding to synthetic membrane vesicles, membrane bilayer insertion rates, and ion channel activity. Finally, we probe and characterize the helix-to-coil transition with steady-state and time-resolved fluorescence spectroscopy.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cytotoxicity Assay of Colicin E1The cytotoxic activity of WT colicin E1 and mutant proteins was confirmed through a "spot test" technique using E. coli strain B cells as described previously (17, 18). In brief, 5-µl aliquots of serially diluted intact WT or mutant colicin E1 protein samples were applied onto a log phase bacterial lawn of colicin-sensitive E. coli, strain B. Cells were then allowed to grow overnight at 37 °C, and cytotoxic activity was monitored as the clearance of the lawn of sensitive strain B cells. The concentrations of the mutant and the WT proteins, at which clearing zones were observed, were compared as a quantitative measure of the cytotoxicity of each mutant protein relative to WT.
Preparation of VesiclesLUVs were prepared from 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3-[phosphorac-(1-glycerol)] (DOPG) (Avanti Polar Lipids, Alabaster, AL), and N-(trinitrophenyl)phosphatidylethanolamine (TNP-PE) (Sigma, discontinued) in 60:30:10% molar ratio by extrusion through a 100-nm polycarbonate filter (Lipofast, Aventin Inc., Ottawa, Ontario, Canada) in 20 mM DMG, 130 mM NaCl at pH 3.55.0 as described earlier (14, 19). Small unilamellar vesicles were prepared from DOPG and 1-palmitoyl-2-stearoyl(9,10)dibromo-sn-glycero-3-phosphocholine (Avanti Polar Lipids) in a 2:3 molar ratio by sonication in an ultrasonic water bath followed by sequential extrusion through a 100- and 50-nm polycarbonate filter (Avestin Inc., Ottawa, Ontario, Canada) as described earlier (19, 20). The suspension buffer used was 20 mM DMG, 130 mM NaCl, at pH 5.
Asolectin (Sigma) LUVs (20 mg/ml) were loaded with the Cl-sensitive fluorophore, 6-methoxy-N-(3-sulfonopropyl) quinolinium (SPQ, 16 mM) (Molecular Probes, Eugene, OR) and were prepared according to the freeze-thaw technique as described previously (21, 22), and the encapsulation buffer was 100 M KCl, 10 mM DMG, 1 mM CaCl2, 16 mM SPQ at pH 5.0. The phospholipid concentration was determined using the Bartlett assay for phosphorus as described by New (23).
Membrane Binding AssayThe binding of WT and mutant colicin E1 channel peptides to membrane vesicles was assayed through the quenching of Trp fluorescence by TNP-PE vesicles, as previously described (24). Fluorescence measurements were obtained with a PTI Alphascan-2 spectrofluorometer (Photon Technologies International, South Brunswick, NJ) equipped with thermostated cell holders and magnetic stirring. Titration data were recorded at 25 °C, with excitation and emission wavelengths set at 293 and 340 nm, respectively. The spectral bandwidths for the excitation and emission wavelengths were set as 4 and 8 nm, respectively. The fluorescence signals were integrated over a 30-s time window in order to increase the signal-to-noise ratio. Vesicle light scattering was minimized using a 309 nm cut-off filter (Oriel Corporation, Rockford, IL) placed in the emission light path; further light scattering contributions to the signal were measured from the buffer titrations of LUVs under the same conditions as described for the protein-lipid titrations.
Measurement of Membrane Insertion KineticsThe vesicle insertion kinetics for channel proteins were measured by the decrease in the Trp fluorescence of the samples caused by the quenching of the brominated phospholipid (bromine at Cys-9, 10-acyl position) as described earlier (20). The final protein and lipid concentrations used were 0.25 and 125 µM (2:3 (mol/mol) 1-palmitoyl-2-stearoyl(9,10)dibromo-sn-glycero-3-phosphocholine/DOPG)), respectively, in 20 mM DMG, 100 mM NaCl, pH 6.
In Vitro Assay of Channel ActivityVoltage-independent chloride efflux from SPQ-loaded vesicles was measured as described by Illsley and Verkman (25). Dye-loaded vesicles were diluted to 0.3 mg/ml in 20 mM DMG, 100 mM NaNO3 buffer, at the desired pH. The vesicle fluorescence was monitored continuously for 2 min at 20 °C with constant stirring after which protein was added (2 ng/ml, final conc.). Fluorescence measurements were taken using excitation and emission wavelengths of 347 and 445 nm, respectively. The spectral bandwidth for both wavelengths was set to 5 nm. The extravesicular buffer was 100 mM NaNO3, 10 mM DMG, pH 6.0, and the encapsulation buffer was 100 mM KCl, 10 mM DMG, 1 mM CaCl2, at pH 5.0. The total amount of encapsulated Cl was released by the addition of Triton X-100 (0.1%, final concentration) to the sample. The fluorescence changes upon the addition of the protein were reported as the percent maximum SPQ fluorescence (%Fmax); %Fmax(t) = ((F Fb)/(FT Fb)) x 100%, where Fb is the initial residual fluorescence of the dye-loaded vesicles and FT is the maximal fluorescence intensity after detergent lysis of the vesicles. All fluorescence measurements were performed using an Eclipse spectrofluorometer equipped with magnetic stirrer and Peltier thermostated multicell holder (Varian Instruments, Mississauga, Ontario, Canada).
Quantification of Disulfide Bonds within Colicin E1In order to test whether the introduced cysteine residues in the A407C/A411C mutant channel peptide had formed a disulfide bond, the thiol-specific free dye, 2-(4'-maleimidylanilino)naphthalene-6-sulfonic acid (MIANS) was used as a probe for the quantification of the free thiol content of A407C/A411C. First, evidence for disulfide bond formation between Cys-407 and Cys-411 was assayed by spectrophotometrically monitoring the conjugation reaction between free protein sulfhydryl side chains and MIANS. Sample proteins (4 µM) in 100 mM sodium phosphate, 150 mM NaCl, pH 7.5, were reacted with excess MIANS (20 µM) in the absence or presence of a partially denaturing 4 M guanidinium hydrochloride (GdmHCl) solution. The dye/protein reaction mixtures were incubated in the dark and under a nitrogen atmosphere for 1 h. As a control, an equal molar WT channel peptide sample was also reacted with excess MIANS, with or without GdmHCl as described above.
Fluorescence measurements of the protein/dye reaction mixtures were obtained at 25 °C using the PTI spectrofluorometer. The excitation wavelength was set to 322 nm, and emission spectra were scanned from 350 to 550 nm. Both the excitation and emission spectral wavelength bandwidths were set to 4 nm, and the fluorescence spectra were corrected for the appropriate buffer-containing controls.
The number of free sulfhydryl groups of A407C/A411C, under non-denaturing, reducing, and/or partially denaturing conditions was further quantified from the stoichiometry of the MIANS labeling reaction. In brief, the labeling reactions were performed as follows. Purified A407C/A411C in pH 7.5 PBS solutions were treated with a 10-fold molar excess of MIANS for1hinthe dark under a nitrogen atmosphere. 5-Fold molar excess of Tris(2-carboxyethyl)-phosphine hydrochloride (Pierce) and 4 M GdmHCl were used to reduce and then to partially denature the protein, respectively. Unreacted MIANS were separated from the conjugated protein sample using 10-kDa (molecular weight cut-off) MicroconTM concentrators (Amicon, MA), and the concentrated sample was further washed four times with pH 7.5 PBS buffer. The stoichiometry of the protein-MIANS conjugation reaction was calculated as the molar ratio of conjugated MIANS and labeled proteins. The molar concentration of the conjugated MIANS and protein were determined from the absorbance at 322 and 280 nm of the labeled protein using the molar extinction coefficients of 20,877.2 and 28,710 M1 cm1, respectively.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
The pH-dependent binding of WT and "trigger" mutant channel peptides to fluorescently tagged and anionic large unilamellar vesicles (LUVs) was determined according to the method of Heymann et al. (24). This assay exploits the fluorescence resonance energy transfer (FRET) from the Trp residues of the protein to trinitrophenyl (TNP) conjugated to the lipid head group of phosphatidylethanolamine within the membrane (TNP-PE). TNP is an excellent FRET acceptor for excited Trp (donor), having an absorbance spectrum with a maximum at 339 nm, which substantially overlaps the Trp emission spectrum within the range of 320 340 nm (24). The 50% donor-acceptor FRET efficiency distance (Förster distance, R0) for Trp-TNP pairs were reported to be 24 (26) and 27 Å,2 the latter corresponds to the Förster distance specific for colicin E1 Trp FRET to TNP-PE. At a 10% molar ratio TNP-PE lipid concentration within the LUVs, the association of WT colicin E1 with the membrane surface resulted in the quenching of the Trp donor fluorescence of the colicin E1 channel peptide to greater than 95% of the total initial fluorescence (data not shown). This observation was in agreement with earlier findings of Heymann et al. (24) and indicates that at 10% molar ratio concentration in the membrane, the distribution of TNP-PE provides satisfactory coverage of the membrane surface to allow a complete assay of the protein interactions with the membrane.
The pH-dependent membrane-binding profiles of WT and some of the mutant channel peptides prepared for this study are shown in Fig. 2, ad. For the WT colicin E1 channel peptide ([P] = 0.25 µM), the mean values of the binding parameters, Kd and n, at pH 4.0 (Fig. 2, ad) were determined to be 1.5 ± 0.7 nM and 97 ± 2 (mol/mol), respectively, giving an equilibrium association constant (Ka) value of 8.8 ± 2.1 µM1, which is in good agreement with the Ka value reported earlier by Heymann et al. (24). The C-terminal domain of WT colicin E1 shows a strong, cooperative pH dependence for membrane binding with a pKa for association near 4.1 (WT, Fig. 2, ad, diamonds), which agrees well with previous reports (11, 2730). In contrast to the WT protein, the binding isotherms of the double Asp-to-Ser-substituted and disulfide-bonded mutant channel peptides exhibited altered binding profiles with an alkaline-directed shift in pH-dependence (Fig. 2, ad). The single Asp-to-Ser-substituted mutant proteins showed smaller shifts in their pH-dependent binding profiles (data not shown), indicating that the concerted action of multiple acidic residues within the trigger motif is responsible for the pH-activation of colicin E1. The effect on the pH-binding profile for single Asp replacement mutant proteins was evident but marginal (data not shown); however, double replacement mutants, D408S/D423S, D408S/D410S, and D410S/D423S, showed the predicted alkaline pH-shift in their membrane-binding profiles with pKa values between 4.5 and 4.6 (Fig. 2, ac).
|
Characterization of A407C/A411C Disulfide-bonded MutantThe double Cys replacement mutant protein consisting of Ala residues 407 and 411 being substituted with Cys within helix 4 exhibited a similar alkaline pH-shift in membrane binding activity (Fig. 2d). The binding parameters for the A407C/A411C mutant channel peptide in the presence of reducing agents (5 mM DTT, data not shown) did not significantly differ from those observed for the oxidized state (Ka = 1.7 ± 0.3 and 1.4 ± 0.2 µM1 in the absence and presence of DTT, respectively, at pH 4.5; Fig. 2d). The formation of a disulfide bond between Cys-407 and Cys-411 in A407C/A411C was confirmed for the conjugation reaction with the thiol-specific fluorescent probe MIANS, which was used to titrate the free sulfhydryl side chains in this mutant channel peptide. Fig. 3 shows the normalized fluorescence spectra of MIANS with the WT and the A407C/A411C mutant channel peptide under denaturing and non-denaturing conditions. Incubation of excess MIANS with the WT channel peptide (the WT possesses 1 buried Cys residue, Cys-505) under non-denaturing conditions resulted in negligible MIAN conjugation, showing little MIANS fluorescence (Fig. 3, trace 1). Partial unfolding of the WT colicin E1 peptide with 4 M GdmHCl and exposure of Cys-505 led to a substantial increase in MIANS fluorescence (Fig. 3, trace 2). In comparison, incubation of excess MIANS with the A407C/A411C mutant channel peptide under non-denaturing conditions resulted in only nominal MIANS-protein conjugation (Fig. 3, trace 3). This resistance to conjugation with MIANS indicated that the two surface-exposed Cys residues in A407C/A411C were predominantly involved in a disulfide bond. Furthermore, partial unfolding of A407C/A411C with 4 M GdmHCl enhanced the MIANS labeling to a similar extent as seen for the WT protein (Fig. 3, trace 4), indicating that only the sulfhydryl side chain of Cys-405 was free to react with the label in the disulfide mutant protein (MIANS/protein molar stoichiometry of 1:1). Additionally, pre-incubation of the A407C/A411C mutant protein under reducing and non-denaturing conditions prior to MIANS labeling yielded a conjugation stoichiometry of 2.1:1 (mol/mol, MIANS/protein) as expected. Moreover, under reducing and partially denaturing conditions, the stoichiometry of the MIANS-protein conjugation reaction was 3.2:1 (mol/mol, MIANS/protein), again as expected (data not shown). In summary, these observations verify that the cysteine residues as engineered into the channel peptide were components of a cystine bond within the A407C/A411C mutant protein.
|
Triple Asp Replacement Mutant Channel Peptide, D408S/D410S/D423SUnfortunately, the triple replacement mutant D408S/D410S/D423S could not be produced in sufficient quantities to be included in this study. Sixteen liters of E. coli culture resulted in barely detectable protein quantities with substantial proteolytic fragments, indicating the instability of this mutant protein. In comparison, a 2-liter culture of E. coli produced over 50 mg of WT protein. In the context of protein production and stability, it is noteworthy that A407C/A411C and D408S/D410S required large culture volumes (8 16 liters) in order to isolate significant amounts of protein. Due to the difficulties in isolation and reduced stability of the D408S/D410S mutant protein, it also could not be included in the subsequent experiments.
Measurement of the Membrane Insertion Kinetics of the Channel PeptidesThe "trigger mutants" were then measured for their ability to insert into the membrane bilayer a necessary step in the conversion of these channel-forming proteins from water-soluble proteins to ion-conducting channels (12). The time courses for the insertion kinetics of the WT, two single replacement mutants, D408S and D410S, two double replacement Asp mutants, D408S/D423S and D410S/D423S, and the double Ala replacement mutant, A407C/A411C, are shown in Fig. 4A. The membrane insertion mechanism is a complicated, multistep process that reflects the rate-limiting step for this process (20). The initial slopes of the individual kinetic traces shown in Fig. 4A were best fit by least squares linear regression analysis in order to determine the apparent membrane insertion rates for the various mutant channel proteins. The insertion rates (units in relative F/ms) are as follows: trace 1, WT, 2.02 ± 0.10; trace 2, D408S, 2.44 ± 0.48; trace 3, D410S, 3.91 ± 0.98; trace 4, A407C/A411C (no DTT), 4.39 ± 0.29; A407C/A411C (with DTT, trace not shown), 5.43 ± 0.65; trace 5, D408S/D423S, 8.88 ± 1.28; and trace 6, D410S/D423S, 10.14 ± 1.81. It is clear from these data that all of the mutant channel proteins exhibited greater membrane insertion rates than the WT protein. Although the single Asp replacement mutants were only marginally more active (Fig. 4A, ranging between 1.2- and 2.0-fold increase), the double Asp replacement mutant proteins exhibited 4.55-fold greater membrane insertion rates than the WT protein. Therefore, the membrane insertion rates for the various mutant proteins clearly depict the cumulative effects of each Asp-to-Ser mutation within the trigger motif of the channel domain. Furthermore, this rate enhancement at pH 6.0 correlates with the strength of the helix-stabilizing bond that was disrupted by mutation, as seen in the kinetic traces for the two Asp-to-Ser single mutants, D408S and D410S (Fig. 4A, traces 2 and 3, respectively). In this respect, the D408S mutation, in which a simple H-bond was disrupted, resulted in a marginal increase in the insertion rate. In contrast, the disruption of the salt-bridged H-bond (between Asp-410 and Lys-406) from the D410S mutation (Fig. 1, A and B; Fig. 4A, trace 3) caused a significantly greater membrane insertion rate. This trend was also observed for the cumulative effect of the double Asp-to-Ser mutants, D408S/D423S and D410S/D423S, with the latter mutant protein exhibiting slightly greater insertion rates than the former (Fig. 4A, traces 5 and 6, respectively).
|
The binding of these membrane-active proteins to the bilayer surface is strongly influenced by the electrostatic attraction of the positively charged protein for the negatively charged membrane surface (24, 31, 32). The Asp-to-Ser replacements would be expected to remove a negative charge at each of these sites, which potentially could account for the greater affinity and/or insertion rates for the mutant channel proteins. However, the effect was also seen for the double Ala-to-Cys mutant protein, where there was no net change in the charge of helix 4 (Fig. 2d and Fig. 4A, trace 4). This mutant protein was originally designed with the idea of forming a disulfide bond within helix 4 of the trigger motif in order to strap this helix into a locked position that would be pH-insensitive. However, the replacement of Ala-407 and Ala-411 by Cys residues caused a destabilization of helix 4, which flawed our original design but served a second purpose (to be discussed later).
Determination of Channel Peptide Cl Efflux ActivityThe in vitro channel activity of colicin E1 has been measured using a variety of techniques (21, 33, 34), and the channel activities of the WT and various mutant proteins are shown in Fig. 4B. The entire mutant channel proteins exhibited enhanced channel activity rates as compared with the WT protein at pH 6.0, as reported by an increase in SPQ fluorescence (Fig. 4B and Table I). Table I illustrates the Cl efflux rates for WT and mutant channel peptides at pH 5.0 and 6.0. The WT protein showed a Cl efflux rate increase from 0.30 ± 0.03 to 14.9 ± 0.10 (x102% Fmax/s) upon acidification from pH 6.0 to 5.0 (a 50-fold increase). The replacement of Asp residues within the trigger motif activated the channel peptide at both pH 5.0 and 6.0 values. The replacement of a single Asp (D408S and D410S) within the trigger mechanism caused a 3 4-fold and 8 12-fold activation for pH 5.0 and 6.0, respectively. The two double Asp replacement mutant proteins (D408S/D423S and D410S/D423S) showed 8 11- and 30 36-fold increase in channel rates for pH 5.0 and 6.0, respectively. The double Cys replacement channel peptide in the oxidized, disulfide-bonded state exhibited a 15- and 70-fold rate enhancement in channel activity for pH 5.0 and 6.0, respectively. Upon reduction, the chloride efflux rates decreased to 6- and 37-fold (compared with the WT protein) for pH values 5.0 and 6.0, respectively. The calculated parameter,
'0 (Table I), is a measure of the relative sensitivity of the channel peptides to pH activation, in which the higher the
'0 value the more sensitive the protein is to pH activation of channel activity. As expected, the WT protein showed the greatest pH sensitivity (
'0, 49.7) followed by the single Asp replacement mutant proteins, D408S and D410S (
'0 values of 22.1 and 16.9, respectively). The double Asp replacement proteins were considerably less sensitive to changes in the solution pH than the single Asp mutant proteins, ranging from 10.9 13.5 (
'0 values for D408S/D423S and D410S/D423S, respectively). However, the least pH-sensitive channel peptide was the double Cys mutant protein with the oxidized form of this protein showing somewhat greater channel activity than the reduced protein (Table I). Intriguingly, the effect of the Ala to Cys mutations (in the disulfide-bonded state without DTT; Fig. 4B, trace 4; Table I) caused an unexpectedly large increase in the ability of the channel domain to form a functional channel under near neutral pH conditions, which can partially be explained on the basis of the difference in the helix-forming propensity between Ala and Cys (P
= 1.41 and 0.66, respectively) (35). Nonetheless, these data indicate that the Cys-407 Cys-411 disulfide bond teases the channel domain into a state that greatly favors one of the steps in channel formation that is subsequent to the membrane binding and insertion events (20). This observation was not anticipated but, nonetheless, it is intriguing and warrants further investigation. In summary, the channel activity data corroborated the membrane binding and kinetic insertion data for the Asp-to-Ser mutant proteins suggesting that these mutations also invoked an enhanced ability of the channel domain to form functional ion channels at near neutral solution pH values.
|
Fluorescence Measurements of the Helix-to-Coil Transition Previously, our laboratory used time-resolved and steady-state fluorescence spectroscopy to identify the proposed pH trigger motif within colicin E1 (13). Accordingly, herein we prepared a combination mutant that featured the F413W mutation along with the D408/D423S mutations. This mutant protein was then subjected to the same analysis protocol as employed previously for the identification of the pH trigger mechanism (13). In addition, the fluorescence quantum yields of these single Trp channel peptides have been shown to increase significantly upon acidification (pH 6.0 3.5) (13). The time-resolved and the steady-state fluorescence parameters of Trp-413 were once again examined in order to ascertain the structural ramifications of the Asp-to-Ser substitutions within the pH trigger motif of the channel domain. The time-resolved fluorescence lifetime and quantum yield analysis confirmed that replacement of the Asp residues within the proposed trigger motif shifted the helix-to-coil equilibrium in favor of the coil state at near neutral pH values (see Supplemental Material for full details and results).
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The proposed trigger motif shows a significant degree of sequence conservation within the family of channel-forming colicins (Fig. 2b). The Asp-408 residue is absolutely conserved; Asp-410 is conserved except for colicins Ia and Ib, and Asp-423 is homologous in all the channel-forming colicins except for Ib. The degree of evolutionary convergence for these acidic residues in channel-forming colicins is statistically significant and indicates a biological and structural relevance, given that the average sequence identity within the channel domain of these proteins (38%) is just over the 30% threshold generally accepted for closely related protein sequences. Our inability to express and purify sufficient quantities of the triple Asp-to-Ser mutant protein, D408S/D410S/D423S, gives this argument further credence. Interestingly, the lack of absolute sequence conservation of these Asp residues within the colicin Ia and Ib sequences is not altogether surprising. The channel domains of colicin Ia and Ib share a distant sequence homology with the two major subfamilies of pore-forming colicins, the E (E1, -5, -10, and K) and A (A, B, N, U, and Y) groups. Whereas the sequence identity of the E and A subgroups range between 6294 and 50 82%, respectively, the Ia and Ib sequences share a 40% identity between them and only show 36 and 24% sequence identity with the E and A subgroups, respectively.
Structural comparison of the pH trigger within the crystal structures of a few members of the channel-forming colicins are illustrated in Fig. 5. The helix-loop-helix motif that constitutes this pH trigger mechanism (boldface helices) can be seen as a slightly off-centered lid, shielding the hydrophobic -hairpin (boldface helices). This pH sensor motif shows significant structural conservation within the channel-forming colicins, E1, A, Ia, and N, for which a high resolution structure has been determined (32, 4042). The identified H-bond acceptor:donor pairs involved in the proposed acidic pH trigger also show an equivalent degree of conservation in these structures, except for colicin Ia. For colicin A, the H-bond acceptor-donor pairs are as follows Asp-468:Arg-467; Asp-478:Asp-481 (main chain); Asp-481:Lys-485. For colicin N, acceptor-donor pairs are as follows: Asp-262:Asn-259 and Thr-353; Asp-264:Lys-260 (main chain). In both of these proteins, Asp-466 (colicin A equivalent to Asp-408 of colicin E1) and Asp-277 (colicin N equivalent to Asp-423 of colicin E1) are not involved in an H-bond. Although it is likely that these residues may still form equivalent H-bonds that were not resolved in the crystal structure, their contributions to the proposed helix-to-coil equilibrium of the pH sensor may still be manifested through charge-helix dipole interactions (discussed earlier). For colicin Ia, the only conserved Asp residues within the pH trigger region are Asp-512 (equivalent to Asp-408 of colicin E1) and Asp-527 (equivalent to Asp-423 of colicin E1). The absence of an acidic residue at 514, equivalent to the helix-stabilizing Asp-410 of colicin E1, is manifested within the crystal structure of colicin Ia as the loss of one-half turn at the N termini of helix 4 (Fig. 5). The variability in both the residue composition and the helical length of the trigger motif within colicin Ia indicates a more rudimentary or primitive trigger mechanism, which may explain the alkaline-shifted membrane-binding profile for this protein (95% transition to the membrane-bound state at pH 4.8) and perhaps its homolog, colicin Ib (43).
|
We propose that firing the pH trigger mechanism within colicin E1 initially involves breaking the critical H-bonds (and salt bridge) within helices 4 and 5a, which initiates a cascade of specific structural events that loosen the tertiary structure contacts between and within the three helical layers of the soluble channel peptide (Fig. 5, colicin E1; layer A, H1, H2, and H10; layer B, H8, H9, and H5; layer C, H3, H4, H6, and H7) (32). Such events are expected to include the (partial) disruption of the helical interactions within layer C and the expansion of the large cavity existing between layer C and the hydrophobic helical hairpin (H8 and H9) (32). This speculation is in agreement with the results of an earlier solution NMR study of colicin E1, which showed an increase in overall mobility of hydrophilic side chains accompanied by site-specific conformational changes that alter the resonances of Val-399/Leu-400, Tyr-396, and one of the two histidine residues (His-427 and His-440) that accompany the low pH-activation of colicin E1 (44). Interestingly, all of these residues flank the pH trigger motif and form a cluster of hydrophobic contacts with the core -helical hairpin.
The formation of pH-induced intermediates for membrane-targeting proteins such as anthrax, cholera, diphtheria, and Shiga toxins, as well as pH-sensitive envelope glycoproteins (e.g. influenza virus hemagglutinin) can be linked to specific cellular pathways that these proteins follow during intoxication of cells (4550). In contrast, the in vitro acidic activation of the pore-forming colicins has not yet been unambiguously correlated with the existence of such an in vivo pathway. A key obstacle to the role for low pH activation in the in vivo mechanism of colicins arises from the fact the translocation pathways of these toxins is largely devoid of acidic compartments that may mimic the conditions used for in vitro studies. Previously, van der Goot et al. (51) linked the formation of the activated state and membrane insertion mechanism of colicin A to a local low pH environment found at the surface of negatively charged membranes such as the bacterial cytoplasmic membrane. This group argued that the activated "molten globule" of colicin A, observed from in vitro studies, is induced by this local low pH environment and that the protein is initially oriented onto the surface of the membrane through electrostatic interactions between a ring of positively charged residues on the surface of the protein and the negatively charged lipid interface (32, 40, 51).
More recently, Schendel and Cramer (11) demonstrated that the contact volume provided by this low pH layer, in which only 18% of the protein encounters a reduced pH environment, would be insufficient to affect the massive unfolding events expected to result from bulk solution acidification in which a molten globule intermediate is formed. Remarkably, inspection of the crystal structures of channel-forming colicins reveals that the proposed pH trigger is centered within the highly conserved ring of positively charged residues (Fig. 6) believed to provide the docking charges that allow the initial adsorption of the protein onto the surface of the membrane (11, 24, 31, 32, 40, 51, 52). As shown in Fig. 6A, the positively charged binding surface provided by this corona of residues is presumed to be appropriately oriented onto the negatively charged membrane surface by the long connector helix (helix 1) between the C-terminal channel domain and the central receptor-binding domain, which is still in contact with the outer membrane receptor (Fig. 6B). Therefore, the observation of a pH trigger nested within this corona of residues presents a novel insight into a possible in vivo mechanism of activation of colicins. Based on the data from the present study, it now seems clear that the low pH activation of the channel-forming colicins can be ascribed to site-specific acidification of the proteins (6, 13). Contrary to the aforementioned solvation volume effect, the requirement of acidic pH for in vitro activity of pore-forming colicins may be a manifestation of the low pH environment sensed by the pH trigger as the protein makes initial contact with the cytoplasmic membrane of the target cell. In this context, protonation of the critical acidic residues of the pH trigger at the surface of the cytoplasmic membrane would be expected to provide a concerted effect that provides both the structural flexibility requisite for membrane insertion as well as an enhanced effective surface charge necessary for the initial electrostatic interactions.
|
In summary, the fundamental premise behind the pH trigger hypothesis is consistent with the existence of structural "sensors" on the surface of proteins that mediate pH-dependent protein conformational transitions as has been previously argued (53). A classical example of this type of sensor is revealed by the acid-induced native-to-molten globule transition of apomyoglobulin, which is largely influenced by the breakage, at pH 4, of an H-bond formed between His-24 and His-119 of the protein (54). More specifically, such a molecular switch mechanism bears resemblance to the helix-to-coil transition of residues 105113 during the low pH activation of influenza hemagglutinin, in which the protonation of Asp-109 and Asp-112 disrupts a network of H-bonds formed by the carboxyl side chain of these acidic residues to the main chain amide of an -helix (55). Mutant hemagglutinin proteins that feature substitution of the aforementioned acidic residues are able to fuse to membranes at higher pH values than WT, highlighting the critical importance of specific residues that act as pH sensors for activation of this toxin (for review see Refs. 55 and 56). Recent evidence from studies on diphtheria toxin suggests that the cooperative binding of at least three protons to the T-domain can result in the formation of the membrane-activated toxin species that adds further credence to the idea for a universal pH sensor-trigger mechanism in microbial proteins (48). In conclusion, confirmation along with an in-depth structural correlation of this pH trigger motif currently awaits the determination of high resolution structures of more of these acid-activated microbial proteins along with structure-function investigations into the molecular details of this type of cell entry mechanism for foreign, invading proteins.
![]() |
FOOTNOTES |
---|
The on-line version of this article (available at http://www.jbc.org) contains additional text, Table S1, and Fig. S1.
To whom correspondence should be addressed. Tel.: 519-824-4120 (ext. 3806); Fax: 519-766-1499; E-mail: merrill{at}chembio.uoguelph.ca.
1 The abbreviations used are: WT, wild-type; DTT, dithiothreitol; DOPC, 1,2-dioleoyl-sn-glycero-3-phosphocholine; DOPG, 1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1-glycerol)]; FRET, fluorescence resonance energy transfer; GdmHCl, guanidinium hydrochloride; LUVs, large unilamellar vesicles; MIANS, 2-(4'-maleimidylanilino)naphthalene-6-sulfonic acid; SPQ, 6-methoxy-N-(3-sulfonopropyl)quinolinium; TNP-PE, N-(trinitrophenyl)phosphatidylethanolamine.
2 A. Szabo and J. Brennan, unpublished data.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|