Molecular Mobility of Elastin: Effect of Molecular Architecture

Biomacromolecules 2002, 3, 531-537

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Molecular Mobility of Elastin: Effect of Molecular Architecture Valerie Samouillan,*,† Jany Dandurand,† Colette Lacabanne,† and William Hornebeck‡ Laboratoire de Physique des Polyme` res, Universite´ Paul Sabatier, 31062 Toulouse Cedex 4, France; and IFR53 Biomole´ cules, Faculte´ de Me´ decine, 51095 Reims, France Received December 5, 2001; Revised Manuscript Received January 24, 2002

The thermal and dielectric properties of elastin and two soluble derivatives (κ-elastin and derived elastin peptides from enzymatic elastolysis) were investigated in the freeze-dried state in a wide temperature range (from -180 to +220 °C). The glass transition of these amorphous proteins was studied by differential scanning calorimetry (DSC). The dielectric relaxations of both proteins were followed by thermally stimulated currents (TSC), an isochronal dielectric spectrometry running at variable temperature, analogous to a lowfrequency spectroscopy (10-3-10-2 Hz) and by dynamic dielectric spectroscopy (DDS), performed isothermally with the frequency varying from 10-2 to 3 × 106 Hz. The combination of TSC and DDS experiments and the determination of the activation parameters of the relaxation times inform about the molecular mobility of the proteins, both in the glassy state and in the liquid state. Major differences between the relaxation behavior of elastin and its soluble derivatives have been discussed and correlated with the molecular architecture of the proteins. Introduction Elastin is a major extracellular matrix macromolecule that confers elasticity to tissues such as skin, lung, and aorta.1 The unusual and highly characteristic amino acid composition of this protein accounts for its great hydrophobicity. It contains one-third glycine amino acids2,3 and several lysine derivatives that serve as covalent cross-links between protein monomers.4 Elastin is thus a three-dimensional network with 60-70 amino acids between two cross-linking points. This molecular architecture is determinant for its elastic properties, insolubility and resistance to proteolysis.5 Nevertheless, the mechanism of elasticity of hydrated elastin is not wholly understood, and several models have been developed to explain this unique feature. First, elastin has been viewed as an aggregate of tropoelastin globules,6 next as a random network devoid of any organization,7 and then considered as a regular arrangement of successive β turns forming a β spiral providing “librational” entropic elasticity8 or composed of isolated and dynamic β turns providing classical entropic elasticity.9,10 The aim of this study is to investigate the low-frequency chain dynamics of native elastin in the nanometer range in order to correlate its molecular mobility to its physical structure. In a previous publication,11 TSC studies indicated that the dielectric parameters, which constitute the fingerprint of the biopolymer, did not show any significant variations with the degradation of insoluble elastin by elastases for up to 50% elastolysis. That is in keeping with reports indicating that insoluble elastin and κ-elastin derived peptides possessed similar global conformations.12,13 However, since the microBrownian motions liberated at the glass transition generally * Corresponding author. E-mail: [email protected]. † Laboratoire de Physique des Polyme ` res, Universite´ Paul Sabatier. ‡ IFR53 Biomole ´ cules, Faculte´ de Me´decine.

correspond to 20-50 atoms, we predicted that an important cleavage of the chains could induce major change in the glass transition of the polymer, thus resulting in the alteration of ordered structures within elastin. To address this question, we compared elastin and elastin-derived peptides (κ-elastin, EDPs from PPE hydrolysis). For this purpose, we have adapted the techniques of characterization of synthetic polymers to the study of proteins. Differential scanning calorimetry (DSC) has been first used to reveal first and second-order transitions of the biological macromolecules. Moreover, since proteins are constituted by polar repeating units (CO-CR-NH), the dielectric techniques are particularly sensitive to analyze relaxation phenomena. Dynamic dielectric spectroscopy (DDS) and thermally stimulated currents (TSC) have been used to reach the relaxation map of the proteins and to determine their molecular mobility. Experimental Section 1. Materials. Insoluble elastin was purified from bovine ligament neck by Lansing’s method14 and was shown by amino acid analysis to be free of microfibrillar proteins. It was freeze-dried and powdered into grains with diameter < 130 µm. κ-Elastin was obtained after a KOH solubilization of bovine elastin,15 and has an average molecular weight Mn ) 75 000. Elastin-derived peptides result from the supernatant of the digestion of elastin by porcine pancreatic elastase, as described in a previous work.11 They will be called EDPs from PPE in this paper. 2. Differential Scanning Calorimetry (DSC). The DSC thermograms were recorded with a Perkin-Elmer DSC7 differential scanning calorimeter. Samples (5-10 mg) were sealed into aluminum pans, and empty pans were used as references. Investigations were performed between 30 and 250 °C with 20 °C/min heating rates. This heating rate was chosen to obtain well-defined glass transitions and to explore the 20-250 °C range in a fast experiment reducing the degradation phenomena.

10.1021/bm015655m CCC: $22.00 © 2002 American Chemical Society Published on Web 03/01/2002

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Figure 1. DSC thermograms (successive scans) of dry elastin, dry κ-elastin, and dry EDPs from PPE.

3. Dielectric Analysis. Samples (20-40 mg) were compressed (2 × 108 Pa during 2 min), resulting in disks 1 mm thick and 8 mm in diameter and then gold-metallized on both sides. To obtain the proteins in the dry state, the samples were annealed for 1 h at 110 °C in N2 atmosphere and then kept in a vacuum (1.33 × 10-4 Pa) for 2 h. Thermogravimetric analysis checked this procedure to remove protein-bound water. The different hydration levels were obtained by equilibrating the samples over saturated salt solutions of known relative humidity at 20 °C during 24 h. Dynamic Dielectric Spectroscopy (DDS). Measurements of the complex dielectric permittivity * were performed in a frequency range from 10-2 to 3 × 106 Hz with a Novocontrol BDS 4000 system. The samples were kept between two gold-plated stainless steel electrodes of a parallel capacitor. The frequency scans were performed isothermally following a temperature step of 5 or 10 °C. The experimental limit for the loss factor tan δ was about 10-4. Thermally Stimulated Currents (TSC). Thermally stimulated currents measurements were carried out with a set up developed in our laboratory and previously described.16 Samples were placed between two plate stainless steel electrodes, and the sample cell was flushed and filled with dry He. The TSC relaxation spectra were recorded with a very sensitive electrometer (Keithley 642, 10-16 A accuracy) vs temperature during a linear heating rate (7 °C/min, which represents an ideal compromise between signal resolution and temperature regulation) after a static polarization at a given temperature and freezing-in at liquid nitrogen temperature. In the present study, the best poling conditions resulting in reproducible dipolar relaxations were as follows: polarizing field Ep ) 400 V/mm, polarization time tp ) 2 min, and linear heating rate q ) 7 °C/min.

Results and Discussion 1. Thermal Transitions. The successive DSC thermograms of elastin, EDPs from PPE and κ-elastin are reported in Figure 1. The first thermogram for the three samples performed until 150 °C is not shown; it only exhibits a broad endothermic peak, vanishing on successive scans, and attributed in previous works17 to the evaporation and vaporization of residual adsorbed water; as a matter of fact, freeze-dried samples easily adsorb water under ambient conditions. The only intrinsic transition of the three proteins is a glass transition occurring at Tg ) 203 °C, Tg ) 190 °C and Tg ) 170 °C for elastin, EDPs from PPE and κ-elastin, respectively. A very slight endothermic peak, superimposed to the

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glass transition and associated with the disruption of physical bonds is hardly noticed during the first cross of the glass transition. Complementary studies show that this enthalpic relaxation of elastin can be amplified by annealing 15-20 °C below Tg.18 Different proteins possess a similar thermogram when annealed at a sub-Tg temperature,19,20 what suggests that the structural relaxation is identical for proteins and synthetic polymers. So elastin can be considered as a glassy material with a three-dimensional network architecture. No crystalline zones are observed, contrary to the majority of extracellular proteins that possess a first-order transition corresponding to the unfolding of peculiar structures.21 The lack of first-order transition in elastin is indicative of the absence of long-range order, while shortrange order can exist with defined secondary structures as isolated and dynamic β turns or R helices domains.9,10 The decrease of the glass transition temperature of the two soluble forms of elastins was expected, as the network arrangement is well-known to stiffen the structure. The loss of the three-dimensional architecture creates free end-chains, increasing free volume and decreasing cohesion, resulting in the decrease of the glass transition temperature. The greater fall of Tg is observed with KOH hydrolysis, that induces a more drastic cleavage than PPE. The other important thermal event noticed by thermal analysis is the start of κ-elastin degradation above 200 °C, in contrast to insoluble elastin and EDPs from PPE that remain thermally stable in this zone. Complementary TGA analysis17 showed that degradation of insoluble elastin occurs at 285 °C. This observation corroborates previous explanations, as the network arrangement stabilizes the protein. 2. Dielectric Relaxations. Low-Temperature Relaxation Modes (-180 to +40 °C). The real and imaginary parts of the complex permittivity of native elastin, 10% hydrated (initial freeze-dried state, equilibrated under N2), were measured from -140 to +40 °C with a step of 5 °C. Figure 2A shows the variation of ′′ vs frequency and temperature in a three-dimensional plot. The ′′ spectra associated with dielectric loss of elastin show a broad mode in the scanned zone, labeled β mode. The temperature dependence of elastin β mode was analyzed by computing the macroscopic relaxation time τmax ) 1/(2π fmax) related to the maximum of ′′ of each isotherm. By plotting it vs temperature (see Figure 2B), we noted that τmax obeys an Arrhenius law in this temperature zone:

( )

τmax(T) ) τ0 exp

Ea RT

where τ0 is the preexponential factor, Ea is the activation energy, and R is the ideal gas constant. The values of these parameters obtained by a linear regression are reported in Table 1. The same dielectric protocol was applied to κ-elastin; in this case, the low-temperature relaxation map is characterized by a β mode (not shown here). An Arrhenius-like behavior is found again for the variation of τmax(T) (see Figure 2B and Table 1). Both the Arrhenius behavior and the value of Ea, close to 50 kJ mol-1, allow us to ascribe the β mode to a secondary

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Molecular Mobility of Elastin

Figure 2. (A) ′′ spectra of elastin 10% hydrated plotted as a function of frequency for temperatures between -140 and +40 °C. (B) Variations of the relaxation times τmax of the β mode of elastin and κ-elastin from DDS experiments.

Figure 3. (A)Complex TSC spectra of elastin and κ-elastin 5% hydrated recorded in the low-temperature range after a polarization at -20 °C. (B) Evolution of the low-temperature TSC spectra of elastin with hydration (w/w) of the sample. The polarization temperature is equal to -20 °C. Table 1. Activation Parameters of Elastin and κ-Elastin β Mode Computed from a Linear Regression of DDS Experiments

elastin κ-elastin

log τ0

Ea (kJ mol-1)

N

R

-14.3 ( 0.3 -15.1 ( 0.2

45 ( 1 47.5 ( 1

13 16

0.998 17 0.998 61

dipolar relaxation mode, as generally observed in synthetic polymers.22 Moreover, the value of τ0, related to the activation entropy ∆S in the Eyring’s theory by τ0 ) h(kT)-1exp(-∆S/R) (where h is Planck’s constant and k Boltzmann’s constant) is indicative of a slightly cooperative mode (∆S close to zero, corresponding to a value of τ0 close to 10-13 s) according to Starkweather’s criterion.23 Figure 3A shows the complex TSC spectra of elastin, EDPs from PPE, and κ-elastin (5%(0.5%) hydrated, recorded between -180 and 0 °C after a polarization at -20 °C. These low-temperature spectra reveal a broad and welldefined peak (labeled β mode) located between -99 and -94 °C for the three samples. This mode is preceded by a small shoulder at around -160 °C, labeled γ mode, which will not be discussed in this paper as it occurs in the limit of the resolution zone.

Contrary to the DDS cell, the TSC cell allows us to make the moisture level vary: in Figure 3B, we present the variation of the TSC spectra of elastin with hydration. Contrary to the γ mode, the β mode is widely affected by the water content: it is shifted toward lower temperatures and magnified as the hydration level increases. This feature illustrates the plasticizing effect of water on the β relaxation. Moreover, the location of the β mode observed by DDS can be extrapolated to the equivalent frequency of TSC (close to 10-3 Hz with a 7 °C/min heating rate24) by replacing τmax by (1/2π10-3) in the Arrhenius’s law previously computed. This calculus leads to a temperature location equal to -120 ( 10 °C for the β mode, which confirms the existence of a depolarization peak in this zone. The good agreement between DDS prediction at 10-3 Hz and TSC measurements for a given hydration level allows us to assert the common origin of β modes observed by DDS and TSC. According to literature data on a wide class of macromolecules as polyamides,25 polyamines, and proteins,26 the great influence of water on the β mode leads us to associate it with relaxing polar sequences of the polypeptidic chain. The plasticizing effect is due to the replacement of protein/ protein hydrogen bounds by protein/water hydrogen bounds,

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Figure 4. M′′ spectra of dry elastin plotted as a function of frequency for temperatures between 50 and 225 °C.

that increase the mobility of the chains.25 Water molecules as well act as electrostatic screens between polar groups, neutralizing the dipole/dipole interactions between polypeptidic units and polar residues in folded proteins.26 These results confirm the essential role of water in the molecular dynamics of elastin as predicted by the Tamburro’s model: 10,27 undoubtedly, water facilitates the motions of the polypeptide chains, and the hydrated relaxed protein must undergo mainly chaotic, Brownian-like motions, behaving as a fractal system of high entropy. As for the comparison between elastin and its two soluble forms, a similar behavior of the β mode is noticed: a similar shape for TSC spectra and close activation values for DDS experiments. We can conclude that the molecular architecture has no influence on the localized relaxation modes of the protein. Both the physical and chemical environments of the relaxing entities remain the same in all the cases. High-Temperature Relaxation Modes (100-220 °C). The real and imaginary parts of the complex permittivity of elastin were measured from 50 to 225 °C with a step of 10 °C up to 170 °C and with a step of 5 °C up to 225 °C. The ′′ spectra associated with dielectric losses in dry elastin (not shown here) show no peak in the probed region, but only an increase of ′′ with rising temperature,27 corresponding to an important phenomenon of conductivity in the lowfrequency range. In the isochronal plotting vs temperature, a broad step is visible on the ′ curves for the lower frequencies (f < 100 Hz), indicative of a relaxation phenomenon. Nevertheless, since ′ does not return to a plateau at high temperature, due to the ionic part of conductivity, this kind of representation is avoided for the determination of the relaxation parameters. The analysis of data in terms of the reciprocal of * has been preferred here in accordance with several authors:29-31 M* ) (*)-1 ) M′ + iM′′ M* is so the dielectric analogue of the mechanical modulus. On the M′′ spectra of dry elastin, shown in Figure 4, a well-defined peak appears in the low-frequency side for temperatures upper than 180 °C. This peak, labeled R, is shifted toward high frequency with rising temperature.

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Figure 5. Temperature dependence of the macroscopic relaxation time τmax for the R modes of elastin and κ-elastin deduced from DDS experiments.

Figure 6. ′′ spectra of dry κ-elastin plotted as a function of temperature for frequencies between 10-2 and 3 × 106 Hz.

Additional measurements performed with a blocking electrode consisting in a PTFE film of 10 µm have confirmed the existence of such a relaxation mode on the ′′ spectra of dry elastin,18 validating the mathematical treatment of ′ and ′′ spectra. The values of τmax corresponding to the maxima of M′′ were computed and plotted vs reciprocal temperature in Figure 5. In the case of dry κ-elastin, ′ and ′′ were recorded from 20 to 180 °C with a step of 5 °C. The exploration temperature was limited due to the earlier degradation of the soluble form. Contrary to elastin spectra, two slight shoulders labeled R1 and R2, superimposed with the conductivity, are noticed on the ′′ curves 6 plotted in an isochronal diagram (see Figure 6). Nevertheless, the M* formalism was used once again in order to compare the elastin and κ-elastin relaxations. The variations of the relaxation time τmax associated with R1 and R2 modes are superimposed to those of elastin in Figure 5. The temperature variation of the average relaxation time of the R mode of elastin is well fitted by the VogelFulcher-Tammann law: τ(T) ) τ0v exp

(

B T - T∞

)

with τ0v ) 9.2 × 10-8 ( 0.2 × 10-8 s, Β ) 660 ( 100 K, and T∞ ) 150 ( 12 °C, a value close to Tg - 50 °C. According to the free volume theory developed by Cohen

Molecular Mobility of Elastin

Figure 7. Complex TSC spectra of dry elastin, dry EDPs from PPE, and dry κ-elastin recorded in the high-temperature range after a polarization at 170 °C.

and Turnbull,32 the dielectric relaxation mode revealed by DDS can be so attributed to the dielectric manifestation of dry elastin devitrification. T∞ corresponds to the temperature at which free volume begins increasing when the temperature is raising, allowing segmental motions to occur. This Vogellike behavior can be only observed at a sub-Tg temperature, namely in the rubbery state. These results are in good agreement with mechanical studies performed both on dry and hydrated elastin, which led to a Williams-Landel-Ferry dependence for the mechanical relaxation associated with elastin glass transition.33 A distinct behavior is found for κ-elastin: the R1 and R2 modes obey Arrhenius laws, with the following activation parameters: τ0R1 ) 7 × 10-19 s, τ0R2 ) 8 × 10-42 s, ER1 ) 142 kJ mol-1, and ER2 ) 329 kJ mol-1. According to JohariGoldstein’s22 and Starkweather’s23 criteria, these modes are delocalized (large value of the activation energy) and cooperative (τ0 < 10-13 s, and so ∆S is largely superior to 0). At the present time, this behavior is not entirely understood, and the only conclusion that can be drawn is the emergence of two distinct chain dynamics of elastin when disruptions occur in its three-dimensional network. To obtain more information about the delocalized relaxation modes of elastin and κ-elastin, the low-frequency technique of TSC was used: we have reported in Figure 7 the complex spectra of dry elastin and dry κ-elastin recorded after a polarization temperature equal to 170 °C. The TSC spectra of dry EDPs from PPE was superimposed to make easier the comparison. A single, well-defined and reproducible relaxation mode, labeled R mode, 60 times more intense than the β one, is observed at 155 and 145 °C, for elastin and EDPs from PPE, respectively. The TSC global spectrum of κ-elastin is significantly distinct from the elastin and EDPs ones. As in DDS experiments, two modes labeled R1 and R2 have been observed: the R2 mode occurs in the same temperature range as the R mode of elastin whereas the R1 mode is shifted toward low temperature (Tmax ) 116 °C). It is noteworthy that the extrapolation of DDS experiments to 10-3 Hz ends up in the same temperature range for these two modes. The split of the R mode into two modes R1 and R2 for κ-elastin could be associated with some heterogeneity in the chain dynamics of κ-elastin: the drastic but partial hydrolysis

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Figure 8. Variation of the preexponential factor vs the activation energy for the elementary relaxation times of dry elastin (data from ref 11) and dry κ-elastin deduced from the FP method in the hightemperature range.

of insoluble elastin drives to a complex system of linear, both ramified and unramified polypeptidic chains. In the case of EDPs from PPE, the only difference with elastin concerns the shift of R mode toward low temperature, emphasizing the decrease of Tg with hydrolysis because of an internal plasticization by hanging end groups. Nevertheless, in this case, the cleavage of polypeptide chains is not so drastic as to induce heterogeneity of the chain dynamics. The experimental decomposition of the R mode of elastin by the fractional polarization technique (FP) was described in a previous paper.11 A set of Debye spectra was recorded by shifting the polarization window in the R relaxation zone, and the temperature dependence of the extracted relaxation times τi(T) was determined using the Bucci-Fieschi formalism.34 We noticed that the temperature dependence of all the relaxation times isolated by the FP method checks Arrhenius laws, driving a distribution of activation energies Eai and a distribution of preexponential factors τ0i. These distributions of Ea and τ0 associated with the probed distribution of relaxing species allow us to attribute the R mode of elastin to the dielectric manifestation of the glass transition of the protein.11 As seen in Figure 8, wherein are reported the variations of τ0 vs Ea for the R mode of elastin, this main mode corresponds to processes of increasing activation enthalpy (Ea reaching 197 kJ mol-1 at the maximum), and with low value of τ0, corresponding to an important activation entropy (∆S reaching 220 J K-1 mol-1). This peculiar behavior reflects the scanning, by TSC, of dipolar motions of larger and larger magnitude along the backbone (Ea is connected, in the William-HoffmannPassaglia model35 to the length of the relaxing unit), which strongly act on their environment36 (high activation entropy). Moreover, a linear relationship between Ea and log τ0 is found for 13 elementary processes in Figure 8. This relationship can be expressed by the following equation: τ0 ) τc exp

( ) Ea RTc

where τc and Tc are connected to the origin ordinate and to the slope of the straight line, respectively.

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And so the relaxation times τ(T) obeying this relationship can be written as follows: τ(T) ) τc exp

( ( )) Ea 1 1 R T Tc

At the temperature Tc, all the related relaxation times would have the same value τc, which is considered in the literature as a compensation phenomenon.37 In this case, the value of the compensation temperature is Tc ) 195 ( 5 °C. This value, close to the glass transition temperature of dry elastin (Tg ) 203 °C), confirms the association of the R relaxation mode with the dielectric manifestation of the glass transition of the protein. The same FP protocol was applied to κ-elastin, and the values of Ea and τ0 computed from the isolated relaxation times were superimposed in Figure 8. In this case, a linear relationship is detected for 8 elementary processes in the zone of the R1 mode; the value of the slope drives to a compensation temperature Tc ) 170 ( 10 °C. Since the glass transition of κ-elastin is found at 170 °C, we can ascribe the R1 mode of κ-elastin to the dielectric manifestation of the devitrification. The most important difference between the two proteins concerns the distribution of activation energies: as a matter of fact, there is an important restriction of this distribution on the high values side (Ea(max)R1 ) 136 kJ mol-1), corresponding to a reduction of the magnitude of the scanned dipolar motions. This value of Ea(max) is then smaller than the value corresponding to insoluble elastin (Ea(max)R ) 197 kJ mol-1). As Ea(max) is associated with the size of the greater cooperative unit in the WilliamHoffmann-Passaglia model,35 we can deduce that the cooperativity of the delocalized motions along the backbone is restricted in soluble elastin. Conclusion This study confirms that both thermal and dielectric techniques are well suited to get a new insight into the structure/mobility correlations of elastin. On one hand, the β mode corresponding to noncooperative movementsstypically on the order of a few nanometerssis analogous in elastin and its derived peptides, meaning that no influence of architecture is detected at this local level of mobility. Another important result concerns the main relaxation mode of elastin, commonly labeled the R mode. It is noteworthy that the R relaxation process observed for the majority of polymers is brought to the fore by dielectric and mechanical techniques, because it corresponds to the delocalized and cooperative movements along the backbone liberated at the thermal glass transitionstypically some tens of nanometerssresulting in dielectric or mechanical energy losses.29 As a matter of fact, the only difference between of these two techniques lies in the used probe. The R relaxation mode is so largely connected to the macroscopic mechanical properties of macromolecules. In the peculiar case of elastin, this R mode is strongly affected by hydrolysis, as illustrated by the discrepancy between the chain dynamics of elastin and its soluble forms. This allows us to assume

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that the mechanical properties of elastin are deeply conditioned by the chain architecture. The direct observation of the dynamic mechanical behavior of hydrated elastin by dynamic mechanical spectrometry is planned in a very near future. It will be now important to check if the flexibility of the polypeptide chains can be affected under pathological conditions (emphysema, lipid deposition, ...) and if the interaction with cells is modified between elastin and its derived peptides. Moreover, as the mechanical R mode of elastin (and consequently the dielectric R mode) is connected to the viscoelastic behavior of the protein, it will be full of interest to consider elastin in the physiological conditions, namely surrounded by water: in this case, the protein is at the very start of the rubber state33 and so must support the model of entropic elasticity developed by Tamburro et al. References and Notes (1) Linsenmayer, T. F. Cell biology of extracellular matrix, 2nd ed.; Hay, E. D., Ed.; Plenum Press: New York, 1991; pp 4-44. (2) Sandberg, L. B. Int. ReV. Connect. Tissue Res. 1976, 7, 159-210. (3) Yeh, H.; Anderson, N.; Ornstein-Goldstein, N.; Bashir, M. M.; Rosenbloom, J. C.; Abrams, W. R.; Indik, Z.; Yoon, K.; Parks, W.; Mecham, R.; Rosenbloom, J. Biochemistry 1989, 28, 2365-2370. (4) Foster, J. A.; Rubin, L.; Kagan, H. M.; Franzblau, C.; Bruenger, E.; Sandberg, L. B. J. Biol. Chem. 1974, 249, 6191-6196. (5) Mecham, R. P.; Heuser, J. E. Cell biology of extracellular matrix; 2nd ed.; Hay, E. D., Ed.; Plenum Press: New York, 1991; pp 79109. (6) Partridge, S. M. Chemistry and molecular biology of the extracellular matrix; Balazs, E. A., Ed.; Academic Press: London, 1969; Vol. 1, pp 593-616. (7) Hoeve, C. A. J.; Flory, P. J. Biopolymers 1974, 13, 677-686. (8) Urry, D. W. Ultrastruct. Pathol. 1983, 4, 227-251. (9) Tamburro, A. M.; Guantieri, V.; Daga Gordini, D. J. Biomol. Struct. Dyn. 1992, 10, 441-454. (10) Villani, V.; D’Alessio, L.; Tamburro, A. M. Elastin and elastic tissue; Tamburro, A. M., Ed.; Congedo Editore: Maratea, Italy, 1996; pp 31-37. (11) Samouillan, V.; Lamure, A.; Maurel, E.; Lacabanne, C.; Hornebeck, W. Biopolymers 2001, 58, 175-185. (12) Debelle, L.; Alix, A. J. P.; Jacob, M. P.; Huvenne, J. P.; Berjot, M.; Sombret, B.; Legrand, P. J. Biol. Chem. 1995, 270, 26099-26103. (13) Debelle, L. Thesis. The University of Reims Champagne Ardenne, S. N. 950384, Reims, France, 1995. (14) Lansing, A. I.; Rosenthel, T. B.; Alex, M.; Dempsey, E. W. Anat. Rec. 1952, 114, 555-570. (15) Jacob, M. P.; Hornebeck, W. Front. Matrix Biol. 1985, 10, 92-129. (16) Teyssedre, G.; Mezghani, S.; Bernes, A.; Lacabanne, C. Thermally stimulated currents of polymers, In Dielectric spectroscopy of polymeric materials. Fundamental and applications; Runt J. P, Fitzgerald J. J., Eds.; American Chemical Society: Washington, DC, 1997; pp 227-255. (17) Samouillan, V.; Dandurand-Lods, J.; Lamure, A.; Maurel, E.; Lacabanne, C.; Gerosa, G.; Venturini, A.; Spina, M. J. Biomed. Mater. Res. 1999, 46, 531-538. (18) Samouillan, V. Thesis. Paul Sabatier University, S. N. 3455, Toulouse, France, 1999. (19) Tseretelli, G.; Smirnova, O. I. Biophysics 1990, 35, 209-214. (20) Huson, M. G. Polym. Int. 1991, 26, 157-161. (21) Walton, A. G.; Blackwell, J. Biopolymers; Academic Press: New York and London, 1973. (22) Johari, G. P.; Goldstein, M. J. Chem. Phys. 1970, 53, 2372-2388. (23) Starkweather, H. W. Macromolecules 1990, 23, 328-332. (24) Van Turnhout, J. Thermally stimulated discharge of polymer electrets; Elsevier Scientific Publishing Co.: Amsterdam, 1975; pp 83-96. (25) Jaffe, M.; Menczel, J. D.; Bessey, W. E. Thermal Characterization of Polymeric Materials; Turi, E. A., Ed.; Academic Press: New York, 1997; pp 1767-1954.

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Molecular Mobility of Elastin (26) Bone, S.; Pethig, R. J. Mol. Biol. 1985, 181, 323-326. (27) Villani, V.; Tamburro, A. M.; Zaldivar Comenges, J. M.; J. Chem. Soc., Perkin. Trans. 2 2000, 11, 2177-2184. (28) Samouillan, V.; Lamure, A.; Lacabanne, C. Chem. Phys. 2000, 255, 259-271. (29) McCrum, N. G.; Read, G.; Williams, B. E. Anelastic and dielectric effects in polymeric solids; John Wiley: New York, 1967. (30) Pathmanathan, K.; Johari, G. P. J. Polym. Sci. 1993, 31, 265-271. (31) Ivanov, D. A.; Jonas, A. M. Polymer 1998, 35, 3577-3581.

(32) (33) (34) (35)

Turnbull, D.; Cohen, M. H. J. Chem. Phys. 1961, 34, 120-125. Lillie, M. A.; Gosline, J. M. Biopolymers 1990, 29, 1147-1160. Bucci, C.; Fieschi, R. Phys. ReV. Lett. 1964, 12, 16-19. Hoffmann, J. D.; Williams, G.; Passaglia, E. J. Polym. Sci. 1966, C14, 173-235. (36) Menegotto, J. Thesis. Paul Sabatier University, 1999. (37) Lavergne, C.; Lacabanne, C. IEEE Electr. Insul. Mag. 1993, 9, 5-21.

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