Biodegradable Poly(ethylene succinate) (PES). 2. Crystal Morphology

Biomacromolecules 2000, 1, 713-720

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Biodegradable Poly(ethylene succinate) (PES). 2. Crystal Morphology of Melt-Crystallized Ultrathin Film and Its Change after Enzymatic Degradation Zhihua Gan, Hideki Abe, and Yoshiharu Doi* Polymer Chemistry Laboratory, RIKEN Institute, Hirosawa 2-1, Wako-shi, Saitama 351-0198, Japan Received June 13, 2000; Revised Manuscript Received August 11, 2000

Poly(ethylene succinate) (PES) ultrathin films with an initial thickness of ∼100 nm were prepared by the solution cast method on either cover glass or freshly cleaved mica as the substrate. The ultrathin films were then melt-crystallized at a given temperature for a certain period of time. The surface morphologies of these films on the substrates were observed by an atomic force microscope (AFM) and an optical microscope (OM) under ambient conditions. Two different crystal morphologies having fibril-like structure and flat-on lamellar crystals with a certain width were formed, and their growth mechanisms were discussed in association with previous kinetic data. It has been shown that at a higher crystallization temperature such as 85 °C (smaller degree of undercooling) the crystal aggregates tend to form lozenge-shaped hedrites which evolved from a single crystal. The enzymatic degradation of PES crystals on the ultrathin films was carried out by using a PHB depolymerase from Pseudomonas stutzeri at room temperature. The crystal morphologies before and after enzymatic degradation were examined by AFM. The lamellar crystals were hydrolyzed into many small fragments, and these fragments had the same thickness as that of the lamellar crystals before enzymatic degradation. The analysis of morphological results for PES lamellar crystals has revealed the existence of many defects on the surface of melt-crystallized lamellar crystals. These defects were preferentially attacked by the enzyme molecules. Hydrolysis starts from the chains folding in crystal defect area and proceeds along the lateral edges, i.e., along the direction perpendicular to the folding chain. Introduction Because of the potential applications of biodegradable polymers in biomedical industries and as environmentally friendly materials, biodegradable polymers have been attracting much attention in the last two decades. A fundamental understanding on the relationship between structure, morphology, and properties will have a notable impact on the future design and synthesis of biodegradable polymers with well-defined chemical structure and desired physical properties. Among the properties of biodegradable polymeric materials, the one that is the most important for practical application is its biodegradation property. Biodegradability is determined mainly by the chemical structure of polymer chains, while it is also strongly dependent on the physical morphology and the degradation environment. As for the inherent structure and morphology of a polymer, besides the stereocomposition,1-4 tacticity4-10 and copolymer composition,11-14 crystallinity, which is inevitable during processing, has a great influence on biodegradation. Now it is generally accepted that crystallinity acts as a hindrance to biodegradability. Biodegradation first takes place in the amorphous region where the erosion rate is much higher than that in the crystalline region.15 Biodegradation in crystalline region * Corresponding author and mail address: Yoshiharu Doi, Polymer Chemistry Laboratory, RIKEN Institute, Hirosawa 2-1, Wako-shi, Saitama 351-0198, Japan. Telephone: +81-48-467-9402. Fax: +81-48-4624667. E-mail: [email protected].

has been well studied, and many papers have been published in recent years to interpret the biodegradation behavior and mechanism. Spherulitic crystal is the most common phenomenon in crystalline polymer morphology and its change before and after biodegradation was studied. A mature spherulite is an array of radiating fibrillar-like structures.16 It was found that during the biodegradation process, the interfibrillar amorphous region was eroded first and the radiating fibrillar-like crystals became clear gradually. Subsequently, these fibrillarlike crystals started to degrade. In addition, during the biodegradation process, holes were observed in the center of spherulites and their sizes increased with time, indicating that the center of spherulite was composed of less ordered lamellar which was more easily degraded.12,15,17,18 The atomic force microscopy (AFM) in situ observation on the degradation of melt-crystallized poly(sebacic anhydride) surface in aqueous environment provided dynamic evidence for the first time that the amorphous materials between spherulitic fibrils were preferentially degraded, resulting in the exposure of the fibrillar structures.19,20 From these results on the spherulites, it is quite clear that the biodegradation of polymer chains in the amorphous region is much faster than that in the crystalline region. However, the degradation in crystalline region is still not clear. For example, there are two opinions on whether the rate of degradation was dependent on crystal size.2,15,17 The holes which appear in the center of spherulites

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and increase in size during degradation indicate that even inside a single spherulite there are differences in crystal order. It implies that the crystal order, orientation, and interaction between these crystals play an important role in biodegradation. To elucidate the mechanism of biodegradation for polymer crystals, single crystals have been utilized as an ideal model due to their well-defined chain-folding lamellar structure and relatively regular surface. Early reports on the degradation of single crystals of polysaccharide showed two kinds of degradation ways.21,22 For β(1f4) xylan single crystals, degradation started preferentially from the edge of the single lamellar crystals and then progressed toward their center area.21 On the contrary, the enzymatic degradation of Nigeran single crystals took place on the surface.22 In recent years many papers have been published on the degradation of poly(3-hydroxybutyrate) (PHB) single crystals under the action of PHB depolymerases from different sources.10,23-26 By using transmission electron microscopy (TEM) to observe the morphology of PHB single crystal after degradation by a PHB depolymerase from Pseudomonas lemoignei, Nobes et al.23 have revealed that the lamellar single crystals were attacked by enzymes from edges and splintered along the long axis of crystal into needlelike morphology as degradation proceeded. Iwata et al.24,25 reported the enzymatic degradation of PHB single crystals by extracellullar PHB depolymerases purified from Pseudomonas stutzeri, Comamonas acidoVorans YM1609, and Alcaligenes faecalis T1, respectively, by using TEM and AFM techniques. By using a special immuno-gold labeling technique, the adsorption of PHB depolymerases on the surface of single crystal was visualized. These enzyme molecules distributed unselectively and homogeneously on the chain-folding surface of the single crystals. However the enzymatic degradation took place preferentially at the chain-packing regions of crystal edges rather than the chain-folding surface of the single crystal. GPC and AFM results indicated that the molecular weight and lamellar thickness remained unchanged before and after degradation. Recently Abe et al. reported the morphology and enzymatic degradation of the lamellar crystals on melt-crystallized ultrathin films (about 100 nm in thickness) of PHB and its copolymers.27 Well-grown lamellar crystals and microfibril crystals along the long axis were observed by using TEM and AFM techniques. Enzymatic degradation resulted in a jagged texture on the lamellar crystal front which was similar to the needlelike structure of PHB single crystal after enzymatic degradation. In comparison to single crystals, polymer ultrathin films of about 100 nm are composed of several lamellar crystals and amorphous layers, whose surface is simpler than that of the bulk sample but whose properties possibly represent those of the bulk materials. By changing the crystallization conditions, ultrathin film of polymer provides us samples with various crystal morphologies which are suitable to study the relationship between crystal structure and degradation mechanism. In the previous paper, we studied the crystal growth kinetics of poly(ethylene succinate) (PES).28 PES is a good sample to examine both the influences of crystal morphology

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and enzyme structure on its degradation. Up to now there is almost no report on the morphology of PES lamellar crystal and its change after degradation. Therefore, in this study, we prepared PES ultrathin film with a thickness of about 100 nm. After melt-crystallizing at different temperatures, the crystal morphologies were observed by optical microscopy (OM) and atomic force microscopy (AFM). Subsequently, the change in the lamellar crystal morphology after enzymatic degradation was examined by AFM and the corresponding degradation mechanism was discussed. Experimental Methods Materials. Poly(ethylene succinate) (PES) was used in this study, and its molecular weights were measured by gel permeation chromatography (GPC) at 40 °C, using a Shimadzu 10A GPC system and a 10A refractive index detector with two joint columns of Shodex K-806 and K-802. Chloroform was used as an eluent at a flow rate of 0.8 mL/ min, and a sample concentration of 2.0 mg/mL was used. Polystyrene standards with low polydispersity were used to make a molecular weight calibration curve. On the basis of this, the weight-average (Mw) and number-average (Mn) molecular weights of PES samples were determined to be 39 400 and 21 100, respectively, and the molecular weight distribution (Mw/Mn) was 1.86. Chloroform was purchased from Kanto Chemical Corp. and used as received. Preparation of PES Ultrathin Films. The PES ultrathin films with a thickness of about 100 nm were prepared by solution cast method on cover glasses (18 × 18 mm in dimension). Briefly, the cover glasses were pretreated by washing with methanol to render the glass surface clear. One droplet of PES solution in chloroform (1.0% w/v) was added onto a cover glass and then covered with another one to spread the solution between the two glasses. After that, the two glasses were slid against each other quickly in opposite direction. PES ultrathin films formed on both glass surfaces after the evaporation of chloroform. Subsequently, the films were heated to a melt state at 180 °C for 1 min and then crystallized isothermally at a given temperature for a given period of time. In consideration of the possible influence of substrate on the crystal morphology of PES ultrathin film, mica was also utilized as substrate. By using the same procedures as mentioned above, the PES ultrathin film was prepared on the surface of freshly cleaved mica and then melt-crystallized for the desired length of time. It was found that there were some influences of substrate on the melt-crystallization of PES ultrathin film. At high crystallization temperatures over 85 °C, due to difficulty in nucleation which led to the formation of a large lamellar crystal, it was easier to prepare PES ultrathin films by melt-crystallizing on the smooth surface of mica than on the relatively rough surface of glass. Therefore, in this study, to prepare good melt-crystallized PES ultrathin films at high temperatures, mica was chosen as the preferred substrate. Morphological Observation of Melt-Crystallized PES Ultrathin Film. The crystalline morphologies of PES ultrathin films after melt-crystallizing at different tempera-

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Figure 1. Optical micrographs of PES ultrathin films melt-crystallized at different temperatures (crossed polarizers): (a) 50, (b) 60, (c) 70, and (d) 80 °C.

tures were examined first with an optical microscope (Nikkon Optiphoto-2) under conditions of crossed and uncrossed polarizers at room temperature. The morphological observation of melt-crystallized PES ultrathin film at lamellar crystal level was carried out using an atomic force microscope (AFM) (SPI37000/SPA300, Seiko Instrument Inc.) under ambient conditions. AFM is a superior tool for the direct observation of polymer sample surface without any needs for sample pretreatment. Both contact and tapping modes were applied in this study for the surface scan of PES lamellar crystals. In contact mode, a triangle cantilever made of Si3N4 was utilized, and its length, spring constant and resonance frequency were 200 µm, 0.021 N/m, and 13 kHz, respectively. For the tapping mode, a rectangle cantilever with a spring constant of 13 N/m and a resonance frequency of 131 kHz was utilized. Enzymatic Degradation of PES Ultrathin Film. The enzymatic degradation of PES ultrathin film was performed by using a PHB depolymerase from P. stutzeri at room temperature. A droplet of enzyme solution (1.49 µmol/mL) was added onto the selected area of PES ultrathin film on the substrate, and the degradation was performed for a given time. After the enzymatic degradation, the droplet of the enzyme solution was removed, and the degraded area was washed three times with pure water. The surface was allowed to dry under ambient conditions before observing the degraded area by AFM.

Results and Discussion Morphologies of Melt-Crystallized PES Ultrathin Films. Figure 1 shows the cross-polarized optical micrographs of PES ultrathin films melt-crystallized at 50, 60, 70, and 80 °C. The spherulites existed in the entire area of the ultrathin films and they were fully impinged on each other after sufficient crystallization time. On the basis of the observation of crystal growth, it was found that in comparison with the PES thick film (∼0.05 mm or more in thickness), the crystallization rate of ultrathin film was much slower and the crystal size was much larger. These results suggest that the thickness of PES ultrathin film has great influences on the primary nucleation rate. Sawamura et al.29 have recently reported that the thickness of polymer film influences the crystal growth rate. For example the crystal growth rate of isotactic polystyrene started to decrease when the thickness of polymer film was below 200 nm. In this paper, our work was not extended to include the crystal growth in PES ultrathin film, but focused instead on the morphological study. The spherulites in Figure 1 also show extinction patterns, but these extinction patterns are different from the typical “Maltese Cross” extinction patterns which usually appear in the thick film. The typical “Maltese Cross” extinction patterns have four black areas inside spherulite radiating along 0, 90, 180, and 270°, respectively. In contrast, the spherulites in PES ultrathin film only had two black areas

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at 135 and 315 °C. With increasing crystallization temperature, the size of spherulite increased due to a decrease in the number of nuclei. However the sensitivity of spherulitic size on crystallization temperature for the ultrathin film was much lower than that for the thick film in which the spherulitic size increased greatly with a rise in temperature.28 When the crystallization temperature was 80 °C, the spherulites had a leaflike structure and the extinction pattern was not clear. This suggests that the final spherulite form had developed from single crystals. To obtain more information on the surface morphology of spherulites at the lamellar crystal level and to understand the growth mechanism, atomic force microscopy (AFM) technique was employed. The AFM images of PES ultrathin films melt-crystallized at 50, 60, 70, and 80 °C have been shown in the previous paper.28 Two distinct crystal morphologies can be detected on the surface of spherulites at different range of crystallization temperatures; one is woven or fibril-like structure along the crystal growth direction from the center of spherulite when crystallization temperatures were 50 and 60 °C, while the other is the flat-on lamellar crystal with a certain width when crystallization temperatures were 70 and 80 °C. In the previous paper, we have demonstrated that the two kinds of lamellar crystal morphologies mentioned above are the reflection of two nucleation regimes for PES (regime II and regime III with a transition at ∼71 °C). The difference in nucleation rate and lateral spreading rate at the front of growing crystal in the different regimes causes the difference in the two dimensions of PES lamellar crystals along the direction of crystal growth and tangential direction of crystal growth. The fibril-like structure is composed of stacks of elongated lamellae. As for the flat-on lamellar crystal, its surface was almost parallel to the substrate. In addition, it was noticed that the side edges of flat-on crystals along the growth direction had either zigzag shapes or nodular shapes when crystallization temperatures were 70 and 80 °C, respectively. To interpret this morphology, we carried out a detailed study on the morphology of PES ultrathin film crystallized at 85 °C. Lamellar Crystal Morphology of PES Ultrathin Film Melt-Crystallized at 85 °C. It was found that various lamellar crystal morphologies developed at 85 °C in the ultrathin film, which provides a good sample for us to understand the crystal growth mechanism. Figure 2 shows the optical micrographs of PES ultrathin films after being melt-crystallized at 85 °C for different durations. These pictures were taken under the condition of uncrossed polarizers. Several lozenge-shaped crystal aggregates can be seen in Figure 2a, having the same shape with PES single crystal.26 These crystal aggregates also have four sectors, similar to that of single crystals, which are shown in the schematic diagram (Figure 3). All the lines in the sectors were parallel to the (110) face of the PES single crystal and had the same distance between each other. The crystal aggregates were observed only in ultrathin film after a 1-day crystallization period. For longer crystallization periods such as 2 days or more, the morphology in Figure 2b was the usual and typical one, which is thought to

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Figure 2. Optical micrographs of PES ultrathin films melt-crystallized at 85 °C for different times (uncrossed polarizers). (a) 1 day; (b) 2 days.

Figure 3. Schematics of PES lozenge-shaped crystal aggregate and single crystal.

have evolved from lozenge-shaped crystal aggregates. Figure 2b suggests that the growth rates of crystal aggregates along the direction of crystallographic axis a and b are much faster than those in the four sector areas. Therefore, the four separated areas of line patterns in Figure 2b are the residua of four sectors of the lozenge-shaped crystal aggregates. The above morphology suggests that the lozenge-shaped crystal aggregates grow from the single crystal, and that they keep the single-crystal growth habit for a certain period of time. To prove it, the morphologies of lamellar crystals in crystal aggregates were observed by AFM. Figure 4 shows the AFM deflection images in the sector area of PES lozenge-shaped crystal aggregates. It was found that the parallel “lines” in the sector were composed of truncated, lozenge-shaped spiral single crystals which developed from dislocation. Both left- and right-hand dislocations occurred (see two arrows in Figure 4b), and the truncated face was always normal to the crystallographic axis a. This result indicates that the lozenge-shaped crystal



Figure 5. Height profile of “lines” in sector area of PES lozengeshaped crystal aggregates indicated by the white line of inset AFM image.

Figure 4. AFM deflection images of lamellar crystal morphology of PES ultrathin film melt-crystallized at 85 °C.

aggregates had actually evolved from single crystals. A question that remains to be answered is why there are many parallel “lines” with periodic distance in the sectors. Figure 5 shows the height profile of the parallel “lines” inside the sectors. The average width of the “line” is about 9 µm. There are two particular phenomena. One is the protrusion of the crystals from the ultrathin films, i.e., the “line” has a height of about 300 nm which is several times higher than the thickness of the initial cast film (about 100 nm in thickness). Similar results from other melt-crystallized polymer thin film have been reported by other researchers.30 The second one is the surface planes of the “line” which are not parallel planes to the substrate but are planes inclined at a small angle of about 3° with substrate. We believe that these two phenomena are related to the dislocation mechanism of crystal growth. It has been reported that initial growth of crystal inclined at a small angle was necessary and important for the continuous growth of spiral crystals by dislocation.31 Such mechanism usually results in the formation of small pits in the originating points of spiral crystals for polymer, which can be observed in Figure 4b and Figure 6a. For the PES

Figure 6. AFM deflection images of PES ultrathin film meltcrystallized at 85 °C in the area growing much faster along the crystallographic axis a* and b*. The arrow indicates the crystal growth direction.

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Figure 7. AFM deflection images during the course of enzymatic degradation of PES ultrathin film melt-crystallized at 80 °C by P. stutzeri PHB depolymerase (1.49 µmol/mL in concentration): (a) 0 min; (b) 7.5 min; (c) 15 min; (d) 25 min.

ultrathin film melt-crystallized at 85 °C, the crystal growth shows the habit of a single crystal via dislocation, but the small angle of initially inclined crystals may cause the formation of spiral crystals at a angle degree of ∼3° to the substrate, as shown in Figure 5. The growth process of inclined crystals may result in the protrusion of crystals. If so the process will be limited by the crystallizable PES chains existing inside the films, i.e., related to the thickness of the film, resulting in the formation of parallel “lines” with periodic distance. Our qualitative results indicated that the distance between the parallel “lines” decreased with a decrease in the thickness of PES ultrathin film. Figure 6 shows the AFM images of the radiating line area in Figure 2b along the crystallographic axis a and b in which crystal growth was faster. The arrow shows the crystal growth direction. The lamellar crystal morphology in Figure 6a shows that, even in the radiating lines area of spherulites for PES ultrathin films melt-crystallized at 85 °C, there are several spiral crystals with lozenge-shaped outlines that have developed from dislocation. Noticeably, the crystallographic axis a of the lozenge-shaped single crystal had

the same direction with the spherulite growth direction. Combined with the morphology shown in Figure 4, the results in Figure 6a further suggest that the spherulites in melt-crystallized PES ultrathin films have actually evolved from single crystals, and the spherulitic growth direction is along the crystallographic axis a. Another morphological feature in Figure 6a is the lateral shapes of lamellar crystals, i.e., the sharp zigzag-shaped edges. The zigzag shaped lamellar crystals are much clearer in Figure 6b. It was noted that the zigzag edges of the lamellar crystals were parallel to the two respective (110) faces, which were contiguous with the obtuse angle of the lozenge-shaped single crystal. These results suggest that the growth of a single lamellar crystal is in accordance with the growth habit of single crystal and that several nuclei are formed and develop to compose one lamellar crystal. This mechanism is in agreement with the nucleation behavior in regime II as analyzed from kinetic results, which indicate that PES is in regime II when the crystallization temperatures is 85 °C. Changes in the Morphology of the Lamellar Crystal after Enzymatic Degradation. The enzymatic degradation



Figure 8. AFM deflection images and corresponding thickness profiles of PES ultrathin film melt-crystallized at 85 °C before (a) and after (b) enzymatic degradation for 3 min by P. stutzeri PHB depolymerase (1.49 µmol/mL in concentration).

of PES ultrathin film was carried out at room temperature by using PHB depolymerase from P. stutzeri. Here we used PES ultrathin films melt-crystallized at 80 and 85 °C as research samples because the PES ultrathin films formed wide and large lamellar crystals which can provide us with a clearer picture after enzymatic degradation. Figure 7 shows the AFM images during the enzymatic degradation process for PES ultrathin films melt-crystallized at 80 °C. Before degradation, stacked lamellar crystals with smooth surfaces (Figure 7a) were clearly observed. During the course of enzymatic degradation, the initially smooth surface of lamellar crystals was gradually replaced by nodular morphology. These observations indicated that the lamellar surfaces were degraded by the P. stutzeri depolymerase. To interpret the degradation mechanism, we further investigated the morphologies of PES ultrathin film meltcrystallized at 85 °C before and after enzymatic degradation for 3 min, and the results are shown in Figure 8. It was noticed that the smooth and integrated lamellar crystals were degraded into many small fragments, and that the lateral edges with straight lines of the initial lamellar crystals were also attacked by the enzymes. The corresponding thickness profile is shown in Figure 8b. It can be seen that the fragment had a thickness of about 10 nm, which is almost the same as the crystal thickness before degradation. These results suggest that although the surface of PES lamellar crystal was attacked by enzyme, the subsequent degradation process took place from the lateral edges of the crystals. PHB depolymerases are known to be composed of two functional domains and a linking region.32 One of the functional domains is the catalytic domain, and the other

one is the binding domain which adsorbs onto the substrate surface and facilitates the approach of the catalytic domain toward the substrate. The enzymatic degradation of PHB crystals by PHB depolymerase occurs in two steps. The first step is the adsorption of the binding domain onto the surface of polymer substrate, and the second step is the hydrolysis reaction, which is carried out by the catalytic domain. The adsorption of the binding domain of PHB depolymerase onto the surface of polymer crystals is considered to contribute to an increase in the polymer chain mobility, which then results in the facile hydrolysis of the polymer chains.24 In this study, we used the PHB depolymerase from P. stutzeri which is composed of both the binding and the catalytic domains.32 It is presumed that the enzymatic degradation of PES lamellar crystals by the PHB depolymerase also proceeds via two steps and that the adsorption of PHB depolymerase also contributes to an increase in mobility of the PES chains to facilitate hydrolysis. In lamellar crystals grown under melt-crystallization condition, the packing of folded chains is relatively poor, especially on the chain-folding surface of lamellar crystal. This is due to the relatively high viscosity of the melt and the relatively fast crystallization rate. Therefore, there may be many defects. As shown in Figure 8a, the corresponding thickness profiles of lamellar crystals before enzymatic degradation indicate that the crystal surface is relatively rough, approximately 2-3 nm in roughness. It is generally accepted that polymer lamellar crystals are composed of folded chains, and that the surface of lamellar crystal is the chain-folding surface. It is however difficult to prepare lamellar crystals with regular surfaces composed of only tight loops of sharp adjacentreentry chain folding. Usually, besides the tight loops, there

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are also some loose loops on the surface of lamellar crystals caused by loose adjacent reentry or loose nonadjacent reentry of folding chains. The majority of loose loops may form an amorphous layer on the lamellar crystal surface, resulting in a rough surface, as observed in melt-crystallized PES lamellar crystals (Figure 8a). Such kind of loose loops on the surface of lamellar crystals as well as the folded chains which connect with these loops in the inner part of the lamellar crystals may constitute the defects in lamellar crystals. The mobility of the folding chains that is connected with the loose loops is higher than that of the folding chains of adjacent reentry, and the packing of folding chains around the defects are loose compared to other areas. Therefore, under the action of PHB depolymerase from P. stutzeri, the loose loops and the folding chains linked to the loose loops are hydrolyzed first by the enzyme molecules and are removed from the lamellar crystals. Thereby, the PHB depolymerase first forms “holes” in the defects on the surface of lamellar crystal. After initiating the hydrolysis process from this hole, the enzyme shaves the folding chains one by one from the edge sides of the “hole”, in other words, the enzyme enlarges the “hole” along all four directions normal to the direction of chain folding. This is the reason why PES integrated lamellar crystals are degraded into many small fragments as shown in Figure 8b but the thickness of the fragments was unaffected. Conclusions Two-dimensional spherulites were developed in PES ultrathin film, and the crystal morphology was dependent on the nucleation regimes. The flat-on and wide lamellar crystals formed at high crystallization temperatures above 70 °C and were found to evolve from single crystals. The morphological analysis of PES lamellar crystal after enzymatic degradation by PHB depolymerase from P. stutzeri suggests that the surface defects were favored for the initial attack by the enzyme molecules leading to possible enhancement in the mobility of loose loops and folded PES chains in the defects. At the initial stage of degradation, polymer chains around the defects are hydrolyzed and removed from the lamellar crystals, resulting in the formation of “holes” in the crystal. In addition, the hydrolysis of folding chains proceeds along the direction normal to the folding chains, i.e., the lateral edges of the “hole” in crystals, resulting in the formation of many fragments which have the same thickness with that of the initial lamellar crystals. Acknowledgment. This work was supported by the grant for Ecomolecular Science Research, RIKEN Institute provided by the Science and Technology Agency (STA) of Japan.

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References and Notes (1) Kemnitzer, J. E.; McCarthy, S. P.; Gross, R. A. Macromolecules 1992, 25, 5927. (2) Doi, Y.; Kumagai, Y.; Tanahashi, N.; Mukai, K. In Biodegradable polymers and plastics; Vert, M., Feijen, J.; Albertsson, A., Scott, G., Chiellini, E., Eds.; Spec. Publ. No. 109; Royal Society of Chemistry: Cambridge, England, 1992; pp 139-147. (3) Abe, H.; Doi, Y. Macromolecules 1996, 29, 8683. (4) Abe, H.; Matsubara, I.; Doi, Y.; Hori, Y.; Yamaguchi, A. Macromolecules 1994, 27, 6018. (5) Jesudason, J. J.; Marchessault, R. H.; Saito, T. J. EnViron. Polym. Degrad. 1993, 1, 89. (6) Hocking, P. J.; Marchessault, R. H.; Timmins, M. R.; Scherer, T. M.; Lenz, R. W.; Fuller, R. C. Macromol. Chem. Rapid Commun. 1994, 15, 447. (7) Hocking, P. J.; Timmins, M. R.; Scherer, T. M.; Lenz, R. W.; Fuller, R. C.; Marchessault, R. H. J. Macromol. Sci., Pure Appl. Chem. 1995, A32, 889. (8) Timmins, M. R.; Lenz, R. W.; Hocking, P. J.; Marchessault, R. H.; Fuller, R. C. Macromol. Chem. Phys. 1996, 197, 1193. (9) Hocking, P. J.; Timmins, M. R.; Scherer, T. M.; Fuller, R. C.; Lenz, R. W.; Marchessault, R. H. Degradable Polymers, Recyclings, and Plastics Waste Arrangement. Plas. Eng. (N.Y.) 1995, No. 29, 297302. (10) Hocking, P. J.; Marchessault, R. H.; Timmins, M. R.; Lenz, R. W.; Fuller, R. C. Macromolecules 1996, 29, 2472. (11) Inoue, Y.; Sano, F.; Nakamura, K.; Yoshie, N.; Saito, Y.; Satoh, H.; Mino, T.; Matsuo, T.; Doi, Y. Polym. Int. 1996, 39, 183. (12) Koyama, N.; Doi, Y. Macromolecules 1997, 30, 826. (13) Doi, Y.; Abe, H. Macromol. Symp. 1997, 118, 725. (14) Abe, H.; Doi, Y.; Hori, Y.; Hagiwara, T. Polymer 1997, 39, 59. (15) Kumagai, Y.; Kanesawa, Y.; Doi, Y. Makromol. Chem. 1992, 193, 53. (16) Bassett, D. C. In Principles of Polymer Morphology; Cambridge University Press: Cambridge, England, 1981; p 16. (17) Tomasi, G.; Scandola, M.; Briese B. H.; Jendrossek, D. Macromolecules 1996, 29, 507. (18) Tsuji, H.; Ikada, Y. J. Polym. Sci., Part A: Polym. Chem. 1998, 36, 59. (19) Shakesheff, K. M.; Davies, M. C.; Roberts, C. J.; Tendler, S. J. B.; Shard, A. G.; Domb, A. Langmuir 1994, 10, 4417. (20) Shakesheff, K. M.; Davies, M. C.; Domb, A.; Jackson, D. E.; Roberts, C. J.; Tendler, K. J. B.; Williams, P. M. Macromolecules 1995, 28, 1108. (21) Chanzy, H.; Comtat, J. Biopolymers 1979, 18, 2459. (22) Marchessault, R. H.; Revol, J. F.; Robbitt, F.; Nordin, J. H. Biopolymers 1980, 19, 1069. (23) Nobes, G. A. R.; Marchessault, R. H.; Chanzy, H.; Briese, B. H.; Jendrossek, D. Macromolecules 1996, 29, 8330. (24) Iwata, T.;, Doi, Y.; Kasuya, K.; Inoue, Y. Macromolecules 1997, 30, 833. (25) Iwata, T.; Doi, Y.; Tanaka, T.; Akehata, T.; Shiromo, M.; Teramachi, S. Macromolecules 1997, 30, 5290. (26) Iwata, T.; Doi, Y. Macromol. Chem. Phys. 1999, 200, 2429. (27) Abe, H.; Kikkawa, Y.; Iwata, T.; Aoki, H.; Akehata, T.; Doi, Y. Polymer 2000, 41, 867. (28) Gan, Z.; Abe, H.; Doi, Y. Biomacromolecules 2000, 1, 704. (29) Sawamura, S.; Miyaji, H.; Izumi, K.; Sutton, S. J.; Miyamoto, Y. J. Phys. Soc. Jpn. 1998, 67, 3338. (30) Sutton, S. J.; Izumi, K.; Miyaji, H.; Miyamoto, Y.; Miyashita, S. J. Mater. Sci. 1997, 32, 5621. (31) Frank, F. C. Discuss. Faraday Soc. 1949, 5, 48. (32) Ohura, T.; Kasuya, K.; Doi, Y. Appl. EnViron. Microbiol. 1999, 65, 189.

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