Clustering of large hydrophobes in the hydrophobic core of ... · PDF fileProtein Engineering...

Hydrophobic Clusters Affect Protein Stability

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Clustering of large hydrophobes in the hydrophobic core of two-stranded α-helical

coiled-coils controls protein folding and stability.

Stanley C. Kwok, and Robert S. Hodges*

Dept. of Biochemistry and Molecular Genetics, Univ. of Colorado Health Sciences

Center, Denver, CO 80262, USA and Dept. of Biochemistry, Univ. of Alberta,

Edmonton, AB, T6G 2H7, Canada.

Running Title: Hydrophobic Clusters Affect Protein Stability

S.C.K. was supported by studentships from the National Science and Engineering

Research Council of Canada and Alberta Heritage Foundation for Medical research.

This research was supported in part by the Canadian Institutes of Health Research, the

Protein Engineering Network of Centers of Excellence, the University of Colorado

Health Sciences Center, and the National Institutes of Health (grant number; R01GM

61855) to R.S.H.

*To whom correspondence should be addressed. Tel. 303-315-8837. Fax. 303-315-1153.

E-mail: [email protected]

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Abstract: 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 α-helical structure in benign conditions. P3 with 3 hydrophobic clusters

was 98% α-helical while P2 was only 65% α-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 spectra

and confirmed by differential scanning calorimetry. The results showed that P3

assembled into a stable α-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 (∆Tm ≈ 18 0C and ∆[Urea]1/2 ≈ 1.5M) compared to

P2. These results suggested that hydrophobic clustering significantly stabilized the

coiled-coil structure and may explain how long fibrous proteins like tropomyosin

maintains chain integrity while accommodating polar or charged residues in regions of

the protein hydrophobic core.


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INTRODUCTION

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 (α-helices and β-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 towards 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


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clustering enhance stability via favorable enthalpic, geometric packing and van der Waals

interactions?

The two-stranded α-helical coiled-coil is the simplest protein fold consisting of two

amphipathic α-helices wound around one another forming a left-handed super coil

stabilized by hydrophobic burial (6,7). All coiled-coils share a characteristic heptad

(seven-residue) repeat denoted as (abcdefg)n in which non-polar 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 α-helices are unstable in aqueous medium. This

hydrophobic surface where amphipathic α-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 stability. Here we present the de novo design

and characterization of two α-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 (Fig.

1.). Their biophysical characteristics are compared by circular dichroism spectroscopy

(CD), 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 (t-Boc) chemistry (reviewed in 14). The


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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 0.73 mmol amino groups per gram (Novabiochem, Switzerland).

The following side-chain protecting groups were used: benzyl (Thr, Ser), cyclohexyl

(Asp), 4-methylbenzyl (Cys), trityl (Asn), and tosyl (Arg). A peptide-resin core (1.0

gram of MBHA resin containing 0.6 mmol of peptide chain was swelled and washed

repeatedly with dichloromethane (DCM) and N,N-dimethylforamide (DMF) in a 25 ml

polypropylene solid-phase extraction reservoir. Activation reagent O-benzotriazol-1-yl-

1,1,3,3 tetramethyluronium hexafluorophosphate (HBTU, 0.45 M) was dissolved in

DMF/DCM/ dimethylsulfoxide (DMSO) (85:10:5 v/v/v) and reacted with excess t-Boc-

protected amino acid (1.1 equivalent to mmol of polypeptide chain) and excess

diisopropylethylamine (DIEA, 1.5 equivalent to mmol polypeptide chain bound to resin)

for 5 min. The activated amino acid ester (4.0 equivalent excess compared to mmol of

polypeptide chain bound to resin) was then coupled onto the solid-phase support by

agitation for an hour. Excess un-reacted amino acids were removed by three alternating

washes of DCM and DMF. Cleavage of the t-Boc and side-chain protecting groups and

the subsequent release of the completed peptides from the MBHA resin support was

achieved with hydrogen fluoride (HF) containing scavengers, 10% (v/v) anisole and 1%

(v/v) 1,2 ethanedithiol (EDT), magnetically stirred for 60 minutes in a reaction vessel

with the temperature-controlled at (-4oC) by immersion in a sodium chloride waterbath.

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 (RPC, reviewed in 16)

on a Zorbax semi-preparative SB-C8 column (250 x 9.4 mm I.D., 5 µm particle size,

300Å pore size) by linear AB gradient elution (0.2% increasing acetonitrile/min), where


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eluent A is 0.05% aqueous trifluoroacetic acid (TFA) and eluent B is 0.05% TFA 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

RPC on a Zorbax analytical 300 Å SB-C8 column (150 x 2.1 mm I.D., 5 µm particle size,

300Å 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 homo-dimeric

coiled-coil was obtained by overnight stirring in a 100 mM NH4HCO3 buffer, pH 8.5, and

the desired product was purified by RPC (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

PO4, pH 7.0 (benign buffer) at 4 oC. A 100 µl aliquot was loaded into the 12 mm, Epon

cell (charcoal-filled), and the initial loading concentrations of peptide stock solutions

ranged from 50 – 500 µM in benign buffer. The samples were spun at 20 oC at 20,000,

25,000 and 35,000 rpm for 24 hours 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:

Mbuoy = Μω (1 - νρ) (1)

where Mbuoy is the measured buoyant weight, Μω is the molecular weight in Daltons, ν is

the partial specific volume of the sample and ρ is the density of the buffer solution. The

partial specific volume of the sample and density of the buffer were calculated using

SednTerp (ver. 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 (ver


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1.035, University of Connecticut), a nonlinear least squares algorithm for equilibrium

ultracentrifugation analyses (17).

Circular Dichroism Spectroscopy

Circular dichroism (CD) spectroscopy was performed on a Jasco-810 spectropolarimeter

with constant N2 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 was used. The concentration of peptide stock solutions was determined

by absorbance at 275 nm in 6 M Urea (extinction coefficient, ε = 1420 cm-1·M-1, one

tyrosine per peptide chain). For wavelength scan analysis, a 5 mg/ml stock solution of

each peptide in 100 mM KCl, 50 mM PO4, 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:

[θ ] = θobs* mrw /10 l c (2)

where θobs 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 interval from 195 to 250 nm. The uncertainty in the molar ellipticity values was

+/-300 deg·cm-1·dmol-1. Protein stability measurements were monitored at wavelength

222 nm, indicative of secondary structure of α-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 oC interval at a scan rate of

60 oC per hour. The temperature dependence of the mean residue ellipticity θ was fitted

to obtain fraction of the unfolded state, PU(t), using a non-linear least-squares algorithm

assuming a two-state unfolding reaction with pre-transitional (folded state, θ N(t)) and

post-translational (unfolded state, θ U(t)) baseline corrections (19,20):

θ (t) = [(1 - PU(t)) * θ N(t)] + [PU(t) * θ U(t)] (3)


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where the pre- and post-transitional baselines are assumed to be linearly-dependent on

temperature, and with θ N(0) and θ U(0) as 0 oC intercepts, respectively:

θ N(t) = θ N(0) – m * T, (4)

and

θ U(t) = θ U(0) – m * T. (5)

The calculated fraction of the unfolded state, PU(t), is given by:

PU(t) = exp [(-∆GU(t) /RT)] / [1 + exp (-∆GU(t) /RT)], (6)

where ∆GU(t) is the apparent Gibbs free energy of folding described by the Gibbs-

Helmholtz equation:

∆GU(t) = ∆Ho (1 - T / T m) + ∆Cp (T - T m) - T ln ( T / T m ) (7)

where tm is the temperature midpoint of the thermal transition, ∆Ho 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 oC), 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 (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 equilibrate overnight before scanning and to ensure accuracy, selected data

points were rescanned to ascertain proper buffer equilibration. The data were fitted to a

linear extraction method (LEM) described previously in (15) to determine denaturation

midpoint, slope associated with the transition and the change in free energy associated

with the transition, ∆Gu. A two-state unfolding model was used to derive peptide

stability values from urea denaturation results:

ff = ([θ ]obs-[θ ]u)/([θ ]f-[θ ]u) (8)


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where [θ ]f and [θ ]u represents the mean residue molar ellipticity for the fully folded and

unfolded species, respectively, and the [θ ]obs is the observed molar ellipticity at a given

denaturant concentration. The free energy of unfolding was derived from the equation:

∆GU = RT ln Ku (9)

where Ku is the equilibrium constant of the unfolding process. In the case of disulfide-

bridged peptides, where the unfolding process is concentration independent, Ku can be

simplified as:

Ku = (1- ff)/(ff) (10)

thus,

∆GU = RT ln (1- ff)/(ff). (11)

Estimates of the free energy of unfolding in the absence of denaturant, ∆GU(water) and

slope term m were obtained by linear extrapolation to zero:

∆GU = ∆GU(water) – m[denaturant] (12)

(24,25), 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 µM of coiled-coil dimer, and peptides were dissolved in 100 mM

KCl, 50 mM PO4, pH 7.0 buffer. The sample solutions and buffer were filtered and

degassed under vacuum and stored at 5 oC. Buffer scans were repeated until identical

baselines were achieved. The heating rate was 60 oC per hour and the cooling rate was

90 oC per hour, with the excess heat monitored from 5 to 70 oC. 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.


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RESULTS

Design of the α-helical Coiled-coils with Different Hydrophobic Clustering

The peptides used in this study were modeled on heptad sequences that had strong α-

helical potential and a heptad repeat (gabcdef)n where non-polar residues at positions a

and d facilitates coiled-coil formation. In the design of these hydrophobic clustered

peptides, we took advantage of the features of the successful α-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 eight heptads (56 residues) based on two repeating heptad sequences:

ExEAxKA and KxEAxEG where positions x are 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. Inter-chain and

intra-chain ionic interactions were engineered by placing Lys and Glu at positions b, e


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and g resulting in ionic stabilization due to inter-chain electrostatic attractions (i to i’ + 5

or g to e’) and intra-chain ionic attractions (i to i +3 or i to i + 4). To promote coiled-coil

formation, a C-terminal disulfide bridge, Gly-Gly-Cys-Tyr linker was introduced to

faciliatate 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 (approximately 50%

α-helical), but in the helix-promoting environment of 50% TFE (30), significant helical

structure was induced in both peptides (Table 1). Though the amino acid sequences of

these peptide 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 ≈

200µM (Table 1), 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% α-helical) at room temperature (Fig. 2, Table 1). The P2 coiled-

coil with two hydrophobic clusters was only 65 % folded at 20 oC in benign buffer

though more helicity was induced at 5 oC (Fig. 3, Panel A). P3 coiled-coil showed a

[θ ]222/208 = 1.02, indicative of a fully-folded coiled-coil (32) whereas that of P2 coiled-

coil is less than one (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


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benign condition. In 50% TFE, both the disulfide-bridged peptides showed nearly 100%

helical content, and the [θ ]222/208 ratios for P3 and P2 were respectively, 0.91 and 0.89,

indicative of the single-stranded α-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 2 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 – 500 µM) 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 1). 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 oC, and a corresponding

increase in urea denaturation midpoint of approximately 1.5 M (Fig. 3, Table 2). To


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calculate the free energy difference between these two analogs, the linear extrapolation

method was used to evaluate the chemical denaturation data (Table 2) 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

kcal•mol-1 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 2), and might describe the non-ideal behavior

of the poorly formed secondary structure of P2. The m slope value generally correlates

with the magnitude of changes in the non-polar accessible surface area between the

folded and the unfolded state (33).

Differential Scanning Calorimetry of Coiled-coils

In addition to the CD experiments, differential scanning Calorimetry (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 2-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 two-state behavior of peptide unfolding/folding

required a minimum of three hydrophobic clusters to form a stably folded protein (Fig. 4,

∆Tm ≈ 42 0C). We then compared the thermodynamics parameter ∆H and Tm for P3

obtained from thermal melting and DSC experiments and found them to be in excellent


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agreement (Table 2.). The estimated ∆Cp was 1.62 kcal•mol-1•

oC-1, which equated to

13.5 cal•mol-1•

oC-1 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 cal•mol-1•

oC-1 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 to 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 two-stranded coiled-coil with a co-operative unfolding transition. In

contrast, P2 with two hydrophobic clusters is only partially folded and significantly less

stable compared to 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 enthalpy component of hydrophobic stabilization, which would affect the

packing of the two interacting α-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, Dũrr and Jelesarov found that the hydrophobic destabilization

of a Leu to Ala substitution is largely entropic at room temperature but becomes

increasingly enthalpic at higher temperature (37).

Historically, hydrophobic cluster analysis (HCA) structural prediction algorithm, based

on the principles that hydrophobic amino acids cluster together in native folded state, has


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been employed to good effect in identifying proteins with little sequence homology, but

with similar overall protein topology (reviewed in 38). Although HCA 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 observed that independent folding subdomains

with different susceptibility to pressure were evident along its length (40). 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 tropomyosin and

indeed found 9 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 hydrophobic core of 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.


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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 is 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. 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 "spring-loaded" 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, 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.


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REFERENCES:

1. Dill, K.A. (1999) Protein Sci 8, 1166-1180.

2. Yao, J., Chung, J., Eliezer, D., Wright, P.E., & Dyson, H.J. (2001) Biochemistry 12,

3561-3571.

3. Colon, W., Elove, G.A., Wakem, L.P., Sherman, F., & Roder, H. (1996) Biochemistry

35, 5538-5549.

4. Shortle, D., & Ackerman, M.S. (2001) Science 293, 487-490.

5. Klein-Seetharaman, J., Oikawa, M., Grimshaw, S.B., Wirmer, J., Duchardt, E., Ueda,

T., Imoto, T., Smith, L.J., Dobson, C.M., & Schwalbe, H. (2002) Science 295, 1719-

1722.

6. Hodges, R.S., Zhou, N.E., Kay, C.M., & Semchuk, P.D. (1990) Pept Res. 3, 123-137.

7. Zhou, N.E., Zhu, B.-Y. & Kay, C.M., & Hodges, R.S. (1992) Biopolymers 32, 419-

426.

8. Lupas, A. (1996) Trends Biochem Sci. 21, 375-382.

9. Wagschal, K., Tripet, B., & Hodges, R.S. (1999a) J. Mol. Biol. 285, 785-803.

10. Wagschal, K., Tripet, B., Lavigne, P., Mant, C.T. & Hodges, R.S. (1999b)

Protein Sci. 11, 2312-2329.

11. Tripet, B., Wagschal, K., Lavigne, P., Mant, C.T., & Hodges, R.S. (2000) J. Mol.

Biol. 300, 377-402.

12. Hodges, R.S., Semchuk, P.D., Taneja, A.K., Kay, C.M., Parker, J.M., & Mant, C.T.

(1988) Pept Res. 1, 19-30.

13. Sereda, T.J., Mant, C.T., Quinn, A.M., & Hodges, R.S. (1993). J Chromatogr. 646,

17-30.


http://ww

w.jbc.org/

Dow

nloaded from

http://www.jbc.org/


18

14. Merrifield, B. (1997) Meth. Enzymol. 289, 3-13.

15. Kwok, S.C., Mant, C.T., & Hodges, R.S. (2002) Protein Sci. 11, 1519-1531.

16. Mant, C.T., Kondejewski, L.H., Cachia, P.J., Monera, O.D., & Hodges, R.S. (1997)

Meth. Enzymol. 289, 426-469.

17. Johnson, M.L., Correia, J.J., Yphantis, D.A., & Halvorson, H.R. (1981) Biophys J.

36, 575-588.

18. Cooper, T.M., & Woody, R.W. (1990) Biopolymers 30, 657-676.

19. Santoro, M.M., & Bolen, D.W. (1988) Biochemistry 21, 8063-8.

20. Lavigne, P., Kondejewski, L.H., Houston, M.E. Jr,, Sonnichsen, F.D., Lix, B., Sykes,

B.D., Hodges, R.S., & Kay, C.M. (1995) J. Mol. Biol. 254, 505-520.

21. Makhatadze, G.I., & Privalov, P.L. (1995). Adv. Protein. Chem. 47, 307-425.

22. Gomez, J., Hilser, V.J., Xie, D., & Freire, E. (1995) Proteins 22, 404-412.

23. Betz, S.F., Liebman, P.A., & DeGrado, W.F. (1997) Biochemistry 36, 2450-2458.

24. Pace, C.N. (1986) Meth. Enzymol. 131, 266-280.

25. Shortle, D. (1989) J. Biol. Chem. 264, 5315-5318.

26. Kellis, J.T. Jr., Nyberg, K., Fersht, A.R. (1989) Biochemistry 28, 4914-4922.

27. Kohn, W.D., Kay, C.M., & Hodges R.S. (1997) J Mol Biol. 267, 1039-1052

28. McClain, D.L., Binfet J.P., & Oakley, M.G. (2001) J Mol Biol. 313, 371-383.

29. O’Shea, E..K., Klemm, J.D., Kim, P.S., & Alber, T. (1991). Science 254, 539-544.

30. Sonnichsen, F.D., Van Eyk, J.E., Hodges, R.S., & Sykes, B.D. (1992) Biochemistry

37, 8790-8798.

31. Zhou, N.E., Kay, C.M., Sykes, B.D., & Hodges, R,S. (1993) Biochemistry 32, 6190-

6197.


http://ww

w.jbc.org/

Dow

nloaded from

http://www.jbc.org/


19

32. Lau, S.Y.M., Taneja, A.K., & Hodges, R.S. (1984) J. Biol. Chem. 317, 129-140.

33. Myers, J.K., Pace, C.N., & Scholtz, J.M. (1995). Prot. Sci. 4, 2138-2148.

34. Privalov, P.L., & Potekhin, S.A. (1986). Meth. Enzymol. 131, 4-51.

35. Makhatadze, G.I., & Privalov, P.L. (1990). J. Mol. Biol. 213 , 385-391.

36. Xie, D., & Freire E. (1994) J. Mol. Biol. 242, 62-80.

37. Dűrr, E., & Jelesarov, I. (2000) Biochemistry 39, 4472-4482.

38. Callebaut, I., Labesse, G., Durand, P., Poupon, A., Canard, L., Chomilier, J.,

Henrissat, B., & Mornon J.P. (1997) Cell Mol Life Sci. 53, 621-645.

39. Sosnick, T.R., Jackson, S., Wilk, R.R., Englander, S.W., & Degrado, W.F. (1996)

Proteins 24, 427-432.

40. Suarez, M.C., Lehrer, S.S., & Silva, J.C. (2001) Biochemistry 40, 1300-1307.

41. Sodek, J., Hodges, R.S., Smillie, L.B., Jurasek, L. (1972) Proc. Natl. Acad. Sci. U.

S. A. 69, 3800-3804.

42. Tripet, B., Van Eyk, J.E., Hodges, R.S. (1997) J Mol Biol 271, 728-750.

43. Carr, C.M., & Kim, P.S. (1993) Cell 73, 823-832.

44. Tripet, B., & Hodges, R.S. (2002) J Struct Biol 137, 220-235.

45. Chana, M., Tripet, B.P., Mant, C.T., Hodges, R.S. (2002) J Struct Biol 137, 206-

219.

46. Thomasin, A.S., Strelkov, S.V., Burkhard, P., Aebi, U., & Parry, D.A.D. (2002) J

Struct Biol 137, 128-145.

47. Araya, E., Berthier, C., Kim, E., Yeung, T., Wang, X., & Helfman, D.M. (2002) J

Struct Biol 137, 176-183.

48. Gee, M.A., & R.B. Vallee. (1998) Eur Biophys J 27, 466-473.


http://ww

w.jbc.org/

Dow

nloaded from

http://www.jbc.org/


20

49. Vallee, R.B., & Gee, M.A. (1998) Trends Cell Biol 8, 490-494

50. Gee, M.A., Heuser, J.E., & Vallee, R.B. (1997) Nature 390, 636-639

51. Gonzalez, L. Jr., Woolfson, D.N., & Alber, T. (1996) Nat Struct Biol 12, 1011-1018.


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Fig. 1. Peptide nomenclature, sequences and schematics of the coiled-coil analogs used

in this study. Top panel shows the sequence of two peptides, P3 and P2 which contain 3

and 2 clusters, respectively. The heptad repeat is denoted as gabcdef which is shown

above the first heptad of the coiled-coil. Bottom panel shows a schematic representation

of the hydrophobic residues in positions a and d where large hydrophobes are shown by

dark circles (Leu, Ile) and small hydrophobes are shown by open circles (Ala). Three

continuous large hydrophobic residues in the core positions d, a, d constitute a

hydrophobic cluster (rectangular box).

Fig. 2. Biophysical characterization of the two-stranded disulfide-bridged coiled-coil

peptides: secondary structural content and oligomeric state. Panel A: Circular dichroism

spectra of cluster peptides in benign condition (50 mM PO4 (K2HPO4/KH2PO4), 100 mM

KCl, pH 7.0 at 20 oC). The concentration of P3 (•) and P2 (o) are 103 and 110 uM,

respectively. Panel B: Sedimentation equilibrium of cluster peptides in benign condition

at 20,000 rpm: the residuals are shown above the UV absorbance data which were best

fitted to a single-species (two-stranded disulfide-bridged coiled-coil model). Initial

peptide concentrations for P3 and P2 were 100 µM.

Fig. 3. Stability comparison of the P3 (3 cluster) and P2 (2 cluster) peptides. Panel A,

temperature melting of the coiled-coils monitored by circular dichroism spectroscopy at

222 nm. Experimental condition was 50mM PO4 (K2HPO4/KH2PO4), 100mM KCl, pH

7.0 buffer at a starting temperature of 5 oC. Panel B, urea denaturation experiments were

carried out at 20°C in a 50mM PO4 (K2HPO4/KH2PO4), 100 mM KCl, pH 7.0 buffer with


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increasing concentrations of urea as denaturant. Peptide P3 (•) and peptide P2 (o) had

respective concentrations of 103 and 110 uM of disulfide-bridged two-stranded coiled-

coil. The ellipticity of peptide P3 was taken as fully-folded to determine fraction folded.

Fig. 4. Stability profile of the 3-clustered P3 peptide: Thermal melt of P3 peptide as a

function of temperature (•) overlaid on the differential scanning calorimetry profile (o).

Experiment condition was 50 mM PO4 (K2HPO4/KH2PO4), 100 mM KCl, pH 7.0 and

peptide concentration was 103 µM of the disulfide-bridged two-stranded peptide.

Fig. 5. Schematic representation of the hydrophobic core residues, in heptad positions a

and d, of the 284-residue coiled-coil protein tropomyosin. Large hydrophobic residues

are shown by dark circles (Leu, Ile, Val, Met, Phe, Tyr) and small nonpolar, 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).


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Table 1 23

P3

P2

Table 1. Biophysical Characterization of the Oxidized and Reduced Hydrophobic Cluster Peptides

PeptideName a

-35,100

-34,600

[θ ]222 (reduced) b

Benign(20° C)

50%TFE

- 34,200

- 22,800

-34,900

-35,200

[θ ]222 (oxidized) c

Benign(20° C)

50%TFE

- 18,800

- 16,500

a Peptides are named by the number of stabilizing hydrophobic clusters shown in Figure 1.b [θ ]222 (reduced) is the mean residue molar ellipticity measured at 222 nm in a reducing buffer 2 mM dithioerythritol (DTE), 50 mM PO4

(K2HPO4/KH2PO4), 100 mM KCl buffer, pH 7.0 in the absence (Benign) or presence of 50% trifluoroethanol (TFE, v/v) at 20 oC. Peptide concentrations were 206 uM and 220 uM (of monomer) for P3 and P2, respectively.

c [θ ]222 (oxidized) is the mean residue molar ellipticity measured at 222 nm in a non-reducing buffer of 50 mM PO4 (K2HPO4/KH2PO4), 100 mM KCl buffer, pH 7.0 in the absence (Benign) or presence of 50% trifluoroethanol (TFE, v/v) at 20 oC. Peptide concentrations were 103 uM and 110 uM (of dimer) for P3 and P2 respectively.

d % Helix was calculated from [θ ]222 based on the benign value divided by the value at 50% TFE multiply by a 100.e The helical ratio [θ ]222/208 was calculated by dividing the observed molar ellipticity value at 222 nm by that at 208 nm.f Apparent molecular weights were determined from sedimentation equilibrium analyses of disulfide-bridged peptides.g Oligomeric state was calculated by dividing the apparent molecular weight by the mass of the disulfide-bridged two-stranded peptide (theoretical MW: 12334).

14,800

13,700

1.2

1.1

Sedimentation Equilibrium

ApparentM.W. f

OligomerRatio g

% Helix (reduced) d

Benign(20° C)

53.6

47.7

% Helix (oxidized) d

Benign(20° C)

98.0

64.8

[θ ] 222/208 e

0.74

0.69

[θ ] 222/208 e

1.02

0.74

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Table 2 24

Table 2. Thermodynamics of Clustered Peptides and Stability Contribution of Hydrophobic Clustering

a Peptides are named by the number of stabilizing hydrophobic clusters shown in Figure 1.b Tm (CD) is the temperature at which there is a 50% decrease in molar ellipticity, [θ ]222, compared to the fully-folded peptide with 3 clusters as determined by circular dichroism spectroscopy.c [Urea]1/2 is the denaturation midpoint of the two-state unfolding of an α-helical coiled-coil to a random coil, i.e., the urea concentration (M) required to achieve a 50% decrease in molar ellipticity, [θ ]222, with a fully-folded coiled-coil taken as –34,200 at 20 oC. d m is the slope term defined by the equation ∆Gu = ∆G(H20) - m [denaturant], and ∆G(H20) is the free energy of denaturation in the absence of denaturant whereas ∆Gu is the free energy of unfolding at a given denaturant concentration.e ∆∆Gu is the free energy difference contributed by a stabilizing cluster: ∆G(H20) (3 Clusters – 2 Clusters).f Tm (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 ∆Cp is the change in heat capacity associated with the the denaturation transition observed in the differential scanning calorimetry experiment.

PeptideName a

Tm (melting) b(oC)

[Urea]1/2c

(M)m, slope d

(kcal•mol-1•M)∆G (H20)

d

(kcal•mol-1)

2.10

0.65

1.35

0.67

2.92

0.86

41.6

23.1

2.1

-

P3

P2

∆∆G (H20)e

(kcal•mol-1)

PeptideName a

42.3

Tm (CD) b(oC)

40.041.6

Tm (DSC) f(oC)

∆H (DSC) h(kcal•mol-1)

∆H (CD) g(kcal•mol-1)

∆Cp (DSC) i(kcal•mol-1 •C-1)

42.9 1.62P3


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Figure 1 25

Amino Acid SequencesPeptideName

P3P2

gabcdefAc-EAEALKA-EIEALKA-KAEAAEG-KAEALEG-KIEALEG-KAEAAEG-KAEALEG-EIEALKA-GGCY-amAc-EAEALKA-EAEALKA-KIEAAEG-KAEALEG-KIEALEG-KAEAAEG-KAEALEG-EIEALKA-GGCY-am

PeptideName

Schematic Representation of Hydrophobic residues at a and d positions

P3P2

a d a d a d a d a d a d a d a d

HydrophobicClusters

3 Clusters2 Clusters


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Figure 2 26

-32,000

-24,000

-16,000

-8,000

205 215 225 235

0

0.30

0.40

0.50

0.60

17.5 18 18.5 20.5 21 21.5

-0.02

0.00

0.02

17.6 18.0 18.4 20.6 21.0 21.4

radius2 / 2 (cm2) radius2 / 2 (cm2)Ab

sorb

ance

R

esid

uals

wavelength (nm)

P2P3

[θ ]

(deg●c

m2 ●

dmol

-1)

A B


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Figure 3 27

Temperature (oC)

Frac

tion

Fold

ed

[Urea] (M)

.

0

0.2

0.4

0.6

0.8

1

10 20 30 40 50 60 70 1 2 3 4

A B


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Figure 4 28

0

500

1000

1500

2000

2500

3000

3500

15 25 35 45 55 65 75

-35,000

-30,000

-25,000

-20,000

-15,000

-10,000

-5,000

0

[θ] 22

2(d

eg●c

m2 ●

dmol

-1)

Temperature (oC)

Cp

(cal●mol -1●

oC-1)


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Figure 5 29

1 99

193

100 190

284


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Stanley C. Kwok and Robert S. Hodgescoiled-coils control protein folding and stability

-helicalαClustering of large hydrophobes in the hydrophobic core of two-stranded

published online July 3, 2003J. Biol. Chem.

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