Clustering of Large Hydrophobes in the Hydrophobic Core of Two-stranded (cid:1) -Helical Coiled-Coils Controls Protein Folding and Stability*

The de novo design and biophysical characterization of two 60-residue peptides that dimerize to fold as parallel coiled-coils with different hydrophobic core clustering is described. Our goal was to investigate whether designing coiled-coils with identical hydrophobicity but with different hydrophobic clustering of non-polar core residues (each contained 6 Leu, 3 Ile, and 7 Ala residues in the hydrophobic core) would affect helical content and protein stability. The disulfide-bridged P3 and P2 differed dramatically in (cid:1) -helical structure in benign conditions. P3 with three hydrophobic clusters was 98% (cid:1) -helical, whereas P2 was only 65% (cid:1) -helical. The stability profiles of these two analogs were compared, and the enthalpy and heat capacity changes upon denaturation were determined by measuring the temperature dependence by circular dichroism spectroscopy and con-firmed by differential scanning calorimetry. The results showed that P3 assembled into a stable (cid:1) -helical two-stranded coiled-coil and exhibited a native protein-like cooperative two-state transition in thermal melting, chemical denaturation, and calorimetry experiments. Although both peptides have identical inherent hydrophobicity (the hydrophobic burial of identical non-polar residues in equivalent heptad coiled-coil positions), we found that the context dependence of an additional hydrophobic cluster dramatically increased stability of P3 ( (cid:2) T m

Understanding protein folding remains a challenging problem: how does information encoded in the amino acid sequence translate into the three-dimensional structure necessary for protein function? Although hydrophobic interactions are generally accepted as the predominant source of free energy change that maintains the folded state, this non-specific stabilization does not describe how the "hydrophobic collapse" guides the formation of specific secondary structure (␣-helices and ␤-sheets) in the final tertiary and quaternary structure in the native protein. The concomitant model suggests that the hydrophobic collapse restricts the conformation of the polypeptide chain into a "molten globule," thus facilitating secondary structure folding in this limited conformational context (1). Examples of hydrophobic interactions participating in the early events of protein folding are observed via stopped flow fluorescence and nuclear magnetic resonance (NMR) studies in apomyogloblin (2) and cytochrome c (3), illustrating the importance of the packing of non-polar residues in stabilizing helix-helix interactions. Recently, non-polar residues have also been observed to form non-native hydrophobic clustering in denatured proteins (4,5), and the authors postulated that non-native hydrophobic interactions can stabilize the long range order of the protein scaffold via an intermediate not observed in the folded state, thus indirectly guiding the extended polypeptide chain toward the correct native fold. Such an observation suggests that the amino acid sequence encodes for structural characteristics other than that of the native fold; in other words, the hydrophobic patterning in the sequence encodes the pathway that ultimately leads to the native functional state. Considering that hydrophobic interactions mediate protein folding both in the folded and the unfolded state, several questions arise: 1) How does a cluster of non-polar residues contribute to stability? 2) Is the free energy derived from the burial of hydrophobic residues simply a sum of the energy derived from the removal of non-polar surface area from aqueous medium? 3) Does hydrophobic clustering enhance stability via favorable enthalpic, geometric packing, and van der Waals interactions?
The two-stranded ␣-helical coiled-coil is the simplest protein fold consisting of two amphipathic ␣-helices wound around one another forming a left-handed supercoil stabilized by hydrophobic burial (6,7). All coiled-coils share a characteristic heptad (7-residue) repeat denoted as (abcdefg) n in which nonpolar residues occupy the a and d positions, forming an amphipathic surface where non-polar interactions allowed assembly of two-, three-, and higher oligomeric states (8). The quantitative contribution of 20 amino acids in positions a and d and their effects on protein stability and oligomerization state have been determined (9 -11). In addition, the secondary structure formation and hydrophobic collapse of coiled-coils are tightly coupled and cooperative since single-stranded amphipathic ␣-helices are unstable in aqueous medium. This hydrophobic surface where amphipathic ␣-helices interact via hydrophobic interactions provides an ideal model to test the effects of hydrophobic clustering. We postulated that hydrophobic clustering in the core of coiled-coils would have a significant influence on secondary structure formation and protein stabil-ity. Here we present the de novo design and characterization of two ␣-helical coiled-coils that have the same inherent hydrophobicity, i.e. the identical hydrophobic core residues (6 Leu, 3 Ile, and 7 Ala residues) but with different clustering of large and small hydrophobic core residues (see Fig. 1.). Their biophysical characteristics are compared by circular dichroism spectroscopy (CD), 1 analytical ultracentrifugation, and differential scanning calorimetry (DSC). The results are discussed in the context of non-polar residue clustering enhancing protein stability.

EXPERIMENTAL PROCEDURES
Peptide Synthesis and Purification-Peptides were synthesized by automated solid-phase methodology described previously (12,13) by conventional t-butyloxycarbonyl chemistry (reviewed in Ref. 14). The peptides were synthesized on an Applied Biosystems model 430A peptide synthesizer as described previously (15). Briefly, the polypeptide chain was assembled on copoly(styrene, 1% divinylbenzene)-4-methylbenzhydrylamine-HCl (MBHA) resin, 100 -200 mesh, substitution of 0.73 mmol amino groups/g (Novabiochem). The following side chainprotecting groups were used: benzyl (Thr, Ser), cyclohexyl (Asp), 4-methylbenzyl (Cys), trityl (Asn), and tosyl (Arg). A peptide resin core (1.0 g of MBHA resin containing 0.6 mmol of peptide chain was swelled and washed repeatedly with dichloromethane and N,N-dimethylformamide in a 25-ml polypropylene solid-phase extraction reservoir. Activation reagent O-benzotriazol-1-yl-1,1,3,3 tetramethyluronium hexafluorophosphate (0.45 M) was dissolved in N,N-dimethylformamide/dichloromethane/dimethyl sulfoxide (Me 2 SO) (85:10:5 v/v/v) and reacted with excess t-butyloxycarbonyl-protected amino acid (1.1 equivalent to mmol of polypeptide chain) and excess diisopropylethylamine (1.5 equivalent to mmol polypeptide chain bound to resin) for 5 min. The activated amino acid ester (4.0 equivalent excess as compared with mmol of polypeptide chain bound to resin) was then coupled onto the solid-phase support by agitation for 1 h. Excess unreacted amino acids were removed by three alternating washes of dichloromethane and N,N-dimethylformamide. Cleavage of the t-butyloxycarbonyl and side chain-protecting groups and the subsequent release of the completed peptides from the MBHA resin support was achieved with hydrogen fluoride containing scavengers, 10% (v/v) anisole, and 1% (v/v) 1,2 ethanedithiol, magnetically stirred for 60 min in a reaction vessel with the temperature controlled at (Ϫ4°C) by immersion in a sodium chloride water bath. The peptide resin was then washed three times with cold diethyl ether to remove scavengers and amino acid-protecting groups. Subsequent resin extraction with glacial acetic acid and overnight lyopholization yielded the crude peptide.
Crude peptides were purified by reversed-phase chromatography (reviewed in Ref. 16) on a Zorbax semipreparative SB-C8 column (250 ϫ 9.4 mm inner diameter, 5-m particle size, 300-Å pore size) by linear AB gradient elution (0.2% increasing acetonitrile/min), where eluent A is 0.05% aqueous trifluoroacetic acid and eluent B is 0.05% trifluoroacetic acid in acetonitrile. The purification was carried out at room temperature with a constant flow rate of 2 ml/min. The purity and homogeneity of the peptide was verified by analytical reversed-phase chromatography on a Zorbax analytical 300-Å SB-C8 column (150 ϫ 2.1 mm inner diameter, 5-m particle size, 300-Å pore size), by quantitative amino acid analysis (Beckman Model 6300 amino acid analyzer), and by electrospray mass spectroscopy on a Fisons Quattro (Fisons, Pointe-Claire, Quebec, Canada). Formation of the disulfide-bridged two-stranded homodimeric coiled-coil was obtained by overnight stirring in a 100 mM NH 4 HCO 3 buffer, pH 8.5, and the desired product was purified by reversed-phase chromatography (described above).
Analytical Ultracentrifugation Equilibrium Experiments-Sedimentation equilibrium analysis was performed on a Beckman XLA analytical ultracentrifuge with absorbance optics at 274 nm for the detection of tyrosine. Samples were first dialyzed exhaustively against an aqueous solution of 100 mM KCl, 50 mM PO 4 , pH 7.0 (benign buffer) at 4°C. A 100-l aliquot was loaded into the 12-mm Epon cell (charcoal-filled), and the initial loading concentrations of peptide stock solutions ranged from 50 to 500 M in benign buffer. The samples were spun at 20°C at 20,000, 25,000, and 35,000 rpm for 24 h to achieve equilibrium, as demonstrated by successive identical radial absorbance scans at 274 nm. The behavior of the peptide species at equilibrium is described by the following equation, where M buoy is the measured buoyant weight, M m is the molecular mass in daltons, is the partial specific volume of the sample, and is the density of the buffer solution. The partial specific volume of the sample and density of the buffer were calculated using SednTerp (version 1.06, University of New Hampshire) using the weighted average of the amino acid content. The peptide oligomerization behavior was determined by fitting the sedimentation equilibrium data from different initial loading concentrations and rotor speeds to various monomer-oligomer equilibrium schemes using WIN NonLIN (version 1.035, University of Connecticut), a non-linear least squares algorithm for equilibrium ultracentrifugation analyses (17). Circular Dichroism Spectroscopy-Circular dichroism (CD) spectroscopy was performed on a Jasco-810 spectropolarimeter with constant N 2 flushing (Jasco Inc., Easton, MD). A Lauda circulating water bath was used to control the temperature of the optic cell chamber, where rectangular cells of 1-mm path length were used. The concentration of peptide stock solutions was determined by absorbance at 275 nm in 6 M urea (extinction coefficient, ⑀ ϭ 1420 cm Ϫ1 ⅐M Ϫ1 , 1 tyrosine per peptide chain). For wavelength scan analysis, a 5 mg/ml stock solution of each peptide in 100 mM KCl, 50 mM PO 4 , pH 7.0 (benign buffer) was diluted and scanned in the presence and absence of 50% trifluoroethanol (TFE). Mean residue molar ellipticity was calculated using the equation, where obs is the observed ellipticity in millidegrees, mrw is the mean residue molecular weight, l is the optical path length of the CD cell (cm), and c is the peptide concentration (mg/ml). Each peptide spectrum was the average of eight wavelengths scans collected at 0.1-nm intervals from 195 to 250 nm. The uncertainty in the molar ellipticity values was Ϯ300 degrees⅐cm 2 ⅐ dmol Ϫ1 . Protein stability measurements were monitored at wavelength 222 nm, indicative of the secondary structure of ␣-helices, by thermal and chemical (urea) denaturations (18).

Temperature-induced Denaturation Monitored by Circular
Dichroism-For thermal melting experiments, data points were taken at 1°C intervals at a scan rate of 60°C/h. The temperature dependence of the mean residue ellipticity was fitted to obtain fraction of the unfolded state, P U(t) , using a non-linear least-squares algorithm assuming a two-state unfolding reaction with pretransitional (folded state, N(t) ) and post-translational (unfolded state, U(t) ) baseline corrections (19,20), where the pre-and post-transitional baselines are assumed to be linearly dependent on temperature, and with N(0) and U(0) as 0°C intercepts, respectively, and The calculated fraction of the unfolded state, P U(t) , is given by, where ⌬G U(t) is the apparent Gibbs free energy of folding described by the Gibbs-Helmholtz equation, where t m is the temperature midpoint of the thermal transition, ⌬H°is the apparent enthalpy of unfolding, and ⌬Cp is the change in heat capacity change associated with protein unfolding. Although ⌬Cp is temperature-dependent (21,22), but in the narrow temperature range of our experiments (5-60°C), this term is generally insensitive to changes (23). These thermodynamic parameters were fitted using the program Igor Pro (WaveMetrics, Inc.) with the protocol described in Ref. 20. Chemical Denaturation Monitored by Circular Dichroism-For chemical denaturation experiments, the stock peptide solution was diluted with appropriate volumes of benign buffer and a stock solution of 10.0 M urea in benign buffer to give a series of data points in increasing denaturant concentration. Data points were left to equili-brate overnight before scanning, and to ensure accuracy, selected data points were rescanned to ascertain proper buffer equilibration. The data were fitted to a linear extrapolation method described previously in Ref. 15 to determine denaturation midpoint, slope associated with the transition, and the change in free energy associated with the transition, ⌬G U . A two-state unfolding model was used to derive peptide stability values from urea denaturation results, where [] f and [] u represents the mean residue molar ellipticity for the fully folded and unfolded species, respectively, and the [] obs is the observed molar ellipticity at a given denaturant concentration. The free energy of unfolding was derived from the equation, where K u is the equilibrium constant of the unfolding process. In the case of disulfide-bridged peptides, where the unfolding process is concentration-independent, K u can be simplified as, thus, Estimates of the free energy of unfolding in the absence of denaturant, ⌬G U(water) and slope term m were obtained by linear extrapolation to zero, where m is the slope associated with unfolding. Differential Scanning Calorimetry-Excess heat versus temperature for the peptides was determined using a Microcal differential scanning calorimeter (Microcal, Northampton, MA). Sample concentrations ranged from 105 to 140 M coiled-coil dimer, and peptides were dissolved in 100 mM KCl, 50 mM PO 4 , pH 7.0, buffer. The sample solutions and buffer were filtered and degassed under vacuum and stored at 5°C. Buffer scans were repeated until identical baselines were achieved. The heating rate was 60°C/h, and the cooling rate was 90°C/h with the excess heat monitored from 5 to 70°C. Each sample was heated and cooled for three cycles to ensure folding reversibility. Data analyses were carried out in Microcal Origin software (Microcal DSC, version 1.2a) using a two-state model with change in heat capacity.

Design of the ␣-Helical Coiled-coils with Different
Hydrophobic Clustering-The peptides used in this study were modeled on heptad sequences that had strong ␣-helical potential and a heptad repeat (gabcdef) n where non-polar residues at positions a and d facilitate coiled-coil formation. In the design of these hydrophobic clustered peptides, we took advantage of the features of the successful ␣-helical coiled-coil models in our laboratory (6, 7, 9 -11), for example, complementary packing in the protein core (26), balance of charged residues across the coiled-coil interface in heptad positions e and g (27,28), and a flexible disulfide bridge linkage (29). The coiled-coil sequences consisted of 8 heptads (56 residues) based on 2 repeating heptad sequences: EXEAXKA and KXEAXEG where positions X represent hydrophobic core positions occupied by Ala, Ile, or Leu in positions a or d (Fig. 1.). We defined a hydrophobic cluster as a consecutive string of three large non-polar residues  (Ile or Leu) in the core positions of the coiled-coil. In our coiled-coils, non-polar residues Leu, Ile, Leu in the consecutive d, a, d heptad positions defined a stabilizing hydrophobic cluster. Our approach was to design two proteins with identical inherent hydrophobicity, i.e. identical number and character of non-polar residues in equivalent coiled-coil core positions but with a different arrangement, i.e. P3 having three hydrophobic clusters and P2 having two (Fig. 1, rectangular boxes). The N-terminal hydrophobic cluster of P3 was disrupted by an interchange of Ile at position 9 and Ala at position 16, both at heptad a positions, to give P2 (Fig. 1, bottom). Therefore, the two analogs have identical inherent hydrophobicity but different clustering patterns. The hypothesis is that the hydrophobic clusters are independent units that contribute to coiled-coil stability and folding when separated along the coiled-coil chain by consecutive strings of Ala residues (Fig. 1, open circles). Thus, this pattern of large and small non-polar core residues helped distinguish the contribution of a hydrophobic cluster from inherent hydrophobicity. Interchain and intrachain ionic interactions were engineered by placing Lys and Glu at positions b, e, and g, resulting in ionic stabilization due to interchain electrostatic attractions (i to iЈ ϩ 5 or g to eЈ) and intrachain ionic attractions (i to i ϩ3 or i to i ϩ 4). To promote coiled-coil formation, a C-terminal disulfide bridge, Gly-Gly-Cys-Tyr linker was introduced to facilitate the formation of a parallel and in-register coiled-coil, and the single Tyr residue allows for protein concentration determination by UV spectroscopy. The disulfide bridge was distant from the N-terminal hydrophobic cluster under investigation (Fig. 1).
Secondary Structure Characterization by Circular Dichroism-Circular dichroism is a sensitive probe of secondary structural features, and this technique was used to detect the difference in helical content between the two peptides. Reduced P3 and P2 peptides were helical in benign condition (ϳ50% ␣-helical), but in the helix-promoting environment of 50% TFE (30), significant helical structure was induced in both peptides (Table I). Although the amino acid sequences of these peptides have a strong underlying helical propensity, potential stabilizing ionic interactions, and amphipathicity, in the reduced state, there is insufficient hydrophobic stabilization to overcome the monomer-dimer equilibrium, at the concentration of ϳ200 M (Table I), to form a fully folded coiled-coil. An interchain disulfide bridge has been shown to enhance coiled-coil folding and stability by eliminating the concentration-dependent monomer-dimer equilibrium that prevented folding (7,31). In contrast to the reduced peptides, both the disulfide-bridged two-stranded coiled-coils P3 and P2 exhibited more helical structure, and the P3 coiled-coil with three hydrophobic clusters was fully folded (98% ␣-helical) at room temperature ( Fig.  2 and Table I). The P2 coiled-coil with two hydrophobic clusters was only 65% folded at 20°C in benign buffer, although more helicity was induced at 5°C (Fig. 3A). P3 coiled-coil showed a [] 222/208 ϭ 1.02, indicative of a fully folded coiled-coil (32), whereas that of P2 coiled-coil is less than 1 (0.74) due to the presence of the single-stranded unfolded state (Fig. 2.). Thus, without the presence of the third hydrophobic cluster, P2 did not fully fold in benign condition. In 50% TFE, both the disulfide-bridged peptides showed nearly 100% helical content, and the [] 222/208 ratios for P3 and P2 were, respectively, 0.91 and 0.89, indicative of the single-stranded ␣-helical conformation (32). The significant helix induction for P2 in TFE, an increase of 35% helicity, showed the underlying high helical propensity of this sequence. However, P2 remained partially unfolded in benign condition because of insufficient hydrophobic stabilization in the hydrophobic core (having only two clusters).
Sedimentation Equilibrium Analyses of Oligomeric States-The packing of hydrophobic core residues had been shown to affect the oligomerization states of coiled-coils (9,10). To determine that P2 and P3 had the same oligomerization state, sedimentation equilibrium analyses were carried out. Using different protein concentrations (from 50 to 500 M) and rotor speeds (20,000 -35,000 rpm), we found that the oligomerization behavior of coiled-coils P3 and P2 were best-fitted to that of a single homogeneous species with molecular weights consistent with a two-stranded coiled-coil ( Fig. 2 and Table I). Overall, sedimentation equilibrium analyses showed that the interchange of hydrophobic residues Ile and Ala that distinguish P2 and P3 did not change the overall oligomerization state, i.e. these coiled-coils are two-stranded and do not assemble into high order oligomerization states. Therefore, we attributed the difference in helicity to a difference in stability due to the presence of an additional stabilizing hydrophobic cluster in peptide P3.
Comparison of Coiled-coil Stability by Thermal and Urea Denaturation-The P3 coiled-coil with three hydrophobic clusters exhibited a highly co-operative unfolding similar to that of native proteins (Fig. 3). In contrast, P2 coiled-coil was only partially folded with marginal stability. We determined the stability of these two analogs using thermal and urea denaturation and found that P3 was significantly more stable than P2, with an increase of thermal midpoint of more than 18°C and a corresponding increase in urea denaturation midpoint of ϳ1.5 M (Fig. 3 and Table II). To calculate the free energy difference between these two analogs, the linear extrapolation method was used to evaluate the chemical denaturation data (Table II) because it is difficult to estimate thermodynamics parameters ⌬H and ⌬Cp from non-ideal thermal melting curves (observed in P2). In addition, a more accurate protocol would be to estimate the free energy change from chemical denaturation because the slope around the urea denaturation transition is reliable (26). The free energy contribution of the N-terminal hydrophobic cluster in the P3 coiled-coil to stability was estimated to be 2.1 kcal⅐mol Ϫ1 per coiled-coil, and this difference in stability can explain why P2 coiled-coil did not fully fold. The slope term m associated with the transition for P3 was significantly higher than that of P2 (Table II)    f t m (DSC) is the temperature midpoint associated with the transition peak during differential scanning calorimetry. g ⌬H (CD) is the change in enthalpy derived from the thermal denaturation experiment using CD spectroscopy.
h ⌬H (DSC) is the change in enthalpy derived from the calorimetry experiment.
i ⌬C p is the change in heat capacity associated with the the denaturation transition observed in the differential scanning calorimetry experiment.
folded and the unfolded state (33).
Differential Scanning Calorimetry of Coiled-coils-In addition to the CD experiments, DSC was also used to measure the contribution of hydrophobic clusters to coiled-coil stability. This technique is one of the most accurate methods for evaluating protein unfolding transitions and analyses of enthalpy and heat capacity changes of temperature-induced processes (34). P3, the three-clustered peptide, showed a cooperative transition with a defined pre-and post-transitional baseline (Fig. 4), but the two-clustered peptide, P2, did not exhibit any measurable transition peak (results not shown), thus illustrating the enthalpic and entropic disruption of the coiled-coil due to the loss of a hydrophobic cluster. Despite the fact that these two coiled-coils have the potential to bury nearly identical hydrophobic surface area, our results suggested that the twostate behavior of peptide unfolding/folding required a minimum of three hydrophobic clusters to form a stably folded protein (Fig. 4, ⌬T m Ϸ 42°C). We then compared the thermodynamics parameter ⌬H and T m for P3 obtained from thermal melting and DSC experiments and found them to be in excellent agreement (Table II.). The estimated ⌬C p was 1.62 kcal⅐mol Ϫ1 ⅐ o C Ϫ1 , which equated to 13.5 cal⅐mol Ϫ1 ⅐ o C Ϫ1 per residue. This value was a relative measurement of the difference in exposed non-polar surface area between the folded and unfolded states (24,36) and agreed well with published values of those of small soluble proteins (35), typically in the range of 8 -15 cal⅐mol Ϫ1 ⅐ o C Ϫ1 per residue. DISCUSSION Hydrophobic interactions contribute significantly to protein stability because the burial of non-polar surface area is thermodynamically favorable in aqueous solution, and this study has shown that the hydrophobic stabilization is context-dependent. In the coiled-coil model, the additional cluster of three large non-polar residues in P3 enhances the folding of secondary structure and protein stability when compared with P2, where this cluster is missing. Both the three-clustered P3 and the two-clustered P2 peptides have the same inherent hydrophobicity (6 Leu, 3 Ile, and 7 Ala residues in the hydrophobic core), yet their folding and stability differ dramatically. P3 with three hydrophobic clusters is a native protein-like twostranded coiled-coil with a co-operative unfolding transition. In contrast, P2 with two hydrophobic clusters is only partially folded and significantly less stable when compared with P3. Furthermore, P3 coiled-coil showed a single unfolding transition in DSC, whereas the unfolding transition was not measurable with P2 under identical experimental conditions. Thus, the disruption of the hydrophobic cluster in P2 drastically decreases the enthalpy component of hydrophobic stabilization, which would affect the packing of the two interacting ␣-helices. The deconvolution of the entropic and the enthalpic components of hydrophobic-residue mutations, i.e. Leu to Ala mutation, is still controversial. For example, Dű rr and Jelesarov (37) found that the hydrophobic destabilization of a Leu to Ala substitution is largely entropic at room temperature but becomes increasingly enthalpic at higher temperature.
Historically, hydrophobic cluster analysis structural prediction algorithm, based on the principles that hydrophobic amino acids cluster together in the native folded state, has been employed to good effect in identifying proteins with little sequence homology but with similar overall protein topology (reviewed in Ref. 38). Although hydrophobic cluster analysis does not predict the hydrophobic effect or protein stability, our results clearly show that significant stabilization can be achieved when hydrophobes cluster in the coiled-coil core. Supporting the results from these prediction programs, experimental data on GCN4 coiled-coil folding suggested that the high energy folding transition state is a hydrophobic collapsed form that contained little secondary structure (39), and therefore, the hydrophobic effect can be an early folding determinant of protein folding, whereas formation of helical secondary structure occurs later.
Hydrophobic clustering may play a significant role in the structure and function of long native coiled-coil proteins. In a recent study on the assembly of tropomyosin, a 284-residue coiled-coil, Silva and co-workers (40) observed that independent folding subdomains with different susceptibility to pressure were evident along its length. When subjected to increase pressure, tropomyosin melted in discrete cooperative blocks along the molecule, and the unfolded domains were likely unstable sites along the coiled-coil chain (40). We examined the hydrophobic clustering in the sequence of rabbit skeletal ␣-tropomyosin and indeed found nine hydrophobic clusters occupied by 3, 4, 5 consecutive large hydrophobes (I, L, M, V, Y, F) in the core a and d positions, separated by destabilizing residues (A, S, T, Q, D, E, and K) in the hydrophobic core (Fig. 5) (41). The authors of that study proposed that increasing osmotic pressure induced water molecules to infiltrate into the less stable "pressure-sensitive" domains in the hydrophobic core of rabbit skeletal ␣-tropomyosin, but the more stable, "pressure-insensitive" domains maintain coiled-coil integrity. It is interesting that our clusters of large hydrophobes (Ile, Leu) and small hydrophobes (Ala) could correspond to the pressure-insensitive and pressure-sensitive domains, respectively. Hydrophobic clustering may be an important mechanism for long native coiled-coil proteins to maintain chain integrity yet still accommodate the burial of polar and charged residues to control stability and facilitate different biological functions. Regardless of the length of the coiled-coil, hydrophobic clusters can serve as "knots" to keep the chain together while allowing flexible regions for function.
Hydrophobic clusters control protein stability and are perhaps an evolutionary feature for native proteins to incorporate structurally important stabilizing regions, as well as less stable and more flexible functional domains in a single protein fold. FIG. 5. Schematic representation of the hydrophobic core residues, in heptad positions a and d, of the 284residue coiled-coil protein tropomyosin. Large hydrophobic residues are shown by dark circles (Leu, Ile, Val, Met, Phe, Tyr), and small non-polar, polar, and charged residues are shown by open circles (Ala, Ser, Thr, Gln, Asp, Glu, and Lys). Three or more continuous large hydrophobic residues in the core positions constitute a hydrophobic cluster (rectangular box). Rabbit skeletal ␣-tropomyosin sequence TPMI_RABIT, P58772.
The destabilizing cluster regions of a coiled-coil may be involved in conformation change that allows for protein-protein interactions. For example, the less stable core regions of tropomyosin could be more easily disrupted for interactions with other proteins (members of the troponin complex and actin involved in muscle regulation) (42). Another example of the importance of flexibility in coiled-coils is the so-called "springloaded" mechanism for viral fusion by the hemagglutinin protein of influenza (43), which allows viral entry into host cells. Conformational change in coiled-coil domains has been implicated, for example, in kinesin, kinesin-like proteins, dynein motor proteins, intermediate filament proteins, and tropomyosins (44 -50). In addition to mediating protein interactions, hydrophobic residues in the coiled-coil core have been known to control oligomerization state in de novo designed coiled-coils (9 -11) and maintain chain orientation in GCN4 by an Asn-Asn pairing (51). Thus, destabilizing clusters could also be involved in these roles. Lastly, hydrophobic clusters are natural nucleation sites for protein folding since they provide the necessary hydrophobic stabilization for early folding intermediates (40). Further investigations into the structural and functional roles of hydrophobic clustering will improve our understanding of the mechanism of coiled-coil folding and the folding of proteins in general.