Cellulose Ester - American Chemical Society

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Biomacromolecules 2009, 10, 2830–2838

Cellulose Ester-graft-poly(ε-caprolactone): Effects of Copolymer Composition and Intercomponent Miscibility on the Enzymatic Hydrolysis Behavior Ryosuke Kusumi, Seung-Hwan Lee, Yoshikuni Teramoto, and Yoshiyuki Nishio* Division of Forest and Biomaterials Science, Graduate School of Agriculture, Kyoto University, Kyoto 606-8502, Japan, and National Institute of Advanced Industrial Science and Technology (AIST), Biomass Technology Research Center, 2-2-2, Hirosuehiro, Kure, Hiroshima 737-0197, Japan Received June 12, 2009; Revised Manuscript Received July 13, 2009

Enzymatic hydrolysis was conducted with Pseudomonas lipase for film samples of graft copolymers of cellulose acetate (CA) and butyrate (CB) with poly(ε-caprolactone) (PCL), CA-g-PCL, and CB-g-PCL, respectively. The two trunk polymers CA and CB, both having the degree of acyl substitution (DS) of >2, are respectively immiscible and miscible with PCL. A hindrance effect of the cellulose ester trunks on the enzymatic attack to the PCL component was observed for the two copolymer series; the situation was more conspicuous in the use of CB trunks. After the selective hydrolytic degradation of the PCL component, a topographical study by AFM revealed that the CA and CB constituents as residues formed a protuberant structure on the surface of the respective film specimens. The altitude and regularity of the protuberances were variant depending on the initial copolymer composition. In a phase-imaging mode of AFM, a hydrolyzed film of CA-g-PCL with an extremely low graftdensity (acetyl DS > 2.95) showed particularly larger CA domains of >25 nm in diameter. The domain sizes were in accordance with a heterogeneity scale in the original intercomponent mixing state estimated by 1H spin-lattice relaxation time (T1H) measurements in solid-state 13C NMR spectroscopy. The present results demonstrate a high potential in application of the PCL-grafted cellulosic copolymers as spatiotemporally biodegradation-controllable materials.

Introduction Today, natural organic resources cellulose and its relatives have been reevaluated as high-potential polymers functionalizable in various chemical ways for many-faceted prospective applications.1,2 Designing of multicompositional systems based on cellulosics, such as polymer blends3-7 and graft copolymers,8-11 is a viable approach not only to improve the original physical properties including processability of the cellulosics but also to expand their availability as highly functionalized polymeric materials.2,12 Applications of the grafting technique to conventional organic esters of cellulose (CEs) can provide a wide range of copolymer compositions, variable depending on the sort of acyl substituent and the degree of substitution (DS) associated with the original three hydroxyl groups as reactive sites on the cellulose backbone. The employment of aliphatic hydroxy acids or cyclic esters as monomer ingredients for the graft reaction onto CEs may be of particular significance, because the resulting graft chains are classified into poly(hydroxyalkanoate)s (PHAs) in a wide sense and the difference in degradability between the CE trunk and PHA graft chains would make it possible to control the overall degradation rate of the polymer material. Recently, the authors’ group has synthesized wide-ranging compositions of cellulose acetate (CA) copolymer with PHA grafts, CA-g-PHAs, by ring-opening polymerization of cyclic ester monomers such as L-lactide (LLA), β-butyrolactone, ε-caprolactone (CL), and so on initiated at the residual hydroxyl positions of CA.8,9 Especially for a CA-g-poly-LLA (CA-g-PLLA) series, Teramoto * To whom correspondence should be addressed. Phone: +81-75-7536250. Fax: +81-75-753-6300. E-mail: [email protected].

et al. demonstrated that the enzymatic degradation behavior of the melt-molded films was widely changeable not only by altering the copolymer composition but also by controlling the supramolecular structure developed in the annealing process.13,14 On the other hand, Kusumi et al. have applied the graft copolymerization of CL to CA and cellulose butyrate (CB) whose acyl DS ranged from 2.1 to more than 2.9;15 the two trunk CEs (CA and CB) are, respectively, immiscible and miscible with PCL.5,6 In that study,15 it was shown for the CE-g-PCL products of PCL-rich compositions that the melt-crystallization kinetics and resulting supramolecular morphology were controllable by adequate selections of the copolymer composition and intercomponent miscibility. Thus, the two CE-g-PCL copolymer series should serve to embody a method of diversifying the degraded-surface morphology as well as the degradation rate of CE/PHA-based materials by counting the difference in miscibility between the polymer pairs employable for the combination. Contemplating the practical functionality of degradable polymers to be used as environmentally conformable and biocompatible materials, for example, in agroindustrial, sanitary, pharmaceutical, and other biorelated fields, we learn that, in addition to some extent of mechanical durability of their moldings, the surface characteristics (e.g., wettability, light reflectance, wear resistance, etc.) are of great significance. In the present study, to investigate a potential of the CE-g-PCL series for such advanced uses as biodegradation-controllable materials, we conducted an enzymatic hydrolysis experiment for different compositions of both CA-g-PCL and CB-g-PCL, by employing Pseudomonas lipase, which shows a strong activity for hydrolysis of PCL.16 The selective hydrolysis behavior was characterized by monitoring the weight loss,

10.1021/bm900666y CCC: $40.75  2009 American Chemical Society Published on Web 08/05/2009

Cellulose Ester-graft-poly(ε-caprolactone)

Biomacromolecules, Vol. 10, No. 10, 2009

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Table 1. Composition Parameters and Molecular Weights of the CE-g-PCL Copolymer Products Used in the Present Study

a

samples

acyl DS

MS

DPs′

WPCL (wt %)

Mwa

Mna

Mw/Mna

CA2.15-g-PCL2.50 CA2.15-g-PCL9.70 CA2.98-g-PCL2.07 CA2.98-g-PCL9.20b CB2.10-g-PCL2.33 CB2.10-g-PCL9.03 CB2.93-g-PCL3.58 CB2.93-g-PCL12.6b

2.15 2.15 2.98 2.98 2.10 2.10 2.93 2.93

2.50 9.70 2.07 9.20 2.33 9.03 3.58 12.6

2.94 11.4 104 460 2.59 10.0 51.1 180

52.9 81.4 45.1 78.5 46.3 77.0 52.6 79.6

226000 384000 1880000 5050000 2090000 3750000 2700000 7070000

124000 191000 743000 2660000 893000 1640000 1340000 3650000

1.82 2.01 2.52 1.90 2.34 2.29 2.02 1.94

Determined by gel permeation chromatography (mobile phase, 0.25 mL/min tetrahydrofuran at 40 °C) with polystyrene standards. b Quoted from ref

15.

compositional change, and surface morphology of film specimens of the graft copolymers, while they were exposed to the enzyme. Discussion of the experimental result took into consideration an effect of the intercomponent miscibility as well as the difference in the initial copolymer composition. A possible heterogeneity in some of the copolymer films, which would be signalized by the enzymatic degradation, was also inspected by 1 H spin-lattice relaxation time measurements in solid-state 13C NMR spectroscopy.

Experimental Section Original Materials and Synthesized CE-g-PCL samples. CA samples of acetyl DS ) 2.15 and 2.98 were kindly supplied by Daicel Chemical Industries, Ltd. and used without further purification. CB samples of butyryl DS ) 2.10 and 2.93 were synthesized with acid chloride/base catalyst from cotton cellulose (Mv ) 252000) via a homogeneous reaction in our laboratory, as has been described in the preceding papers.5,6 Poly(ε-caprolactone) (PCL) with a nominal weightaverage molecular weight of 70000 was provided as PLACCEL H7 from Daicel Chemical Industries, Ltd., and it was used after purification by dissolution in tetrahydrofurane and reprecipitation into distilled water. Lipase from Pseudomonas cepacia (∼50 U/mg) was purchased from Fluka and used as received. Other solvents and chemicals used in this study were purchased from Wako Pure Chemical Industries, Ltd. or Nacalai Tesque, Inc.; those were all of guaranteed reagent grade and used without further purification. CA-g-PCL and CB-g-PCL samples were synthesized by ring-opening graft copolymerization of CL monomer onto CA and CB with a catalyst tin(II) 2-ethylhexanoate (Sn(II)Eht), in a manner similar to that used in previous studies.15 Table 1 summarizes basic data of the copolymer composition parameters, that is, acyl DS, the degree of molar substitution (MS) for the introduced oxycaployl units, the apparent degree of polymerization of the PCL side-chain (DPs′), and the PCL content in weight percent (WPCL), together with data of the average molecular weights (Mw and Mn), for mainly used CA-g-PCL and CB-g-PCL products. A code CEx-g-PCLy denotes CE-g-PCL of acetyl or butyryl DS ) x and oxycaproyl MS ) y. Film Preparation. Films of the two CE-g-PCL series and plain PCL and CEs for enzymatic hydrolysis were molded at 100-260 °C, that is, above the glass transition temperature (Tg) or melting temperature (Tm) of each sample, by using a Toyo-Seiki hot-pressing apparatus with a stainless spacer 0.1 mm thick. For the molding, a pressure was applied to each molten sample gradually to reach 5.0 MPa in 3 min, and subsequently it was increased quickly to 15.0 MPa, followed by maintaining this application for 30 s. Immediately after the pressure was released, the samples were transferred to another compressing apparatus and cold-pressed at 15.0 MPa and 25 °C for 10 min. After released again from the compressed state, strips of circular shape (φ ) 5 mm) were cut off from the molded films and stored at 40 °C in vacuo. However, ungrafted CA2.98 was hardly molded in the manner mentioned above, due to a certain extent of crystallinity as cellulose triacetate. Therefore, a film of CA2.98 was prepared by solution casting with dichloromethane as solvent.

Enzymatic Hydrolysis. Prior to enzymatic hydrolysis, thermal treatments were conducted for the film specimens of CE-g-PCLs and plain PCL and CEs. Each specimen was weighed and heated above Tg or Tm (100-260 °C) in vacuo for 3 min. After the preheating, the sample was immediately transferred to another oven regulated at 25 °C and maintained there for 48 h. A test specimen was placed in a glass tube containing 5 mL phosphate buffer (pH ) 7.0) and 0.5 mg Pseudomonas lipase. The specimen/enzyme incubation was carried out at 37 °C in a rotary shaker (100 rpm). After elapsing of prescribed time, the test tubes each containing a polymer specimen were drawn out of the incubator, and the specimens were taken out and washed thoroughly with distilled water, then dried in vacuo at room temperature until their weights reached constancy. These specimens were evaluated for the time-course of the weight loss in the enzymatic hydrolysis process. Measurements. Differential scanning calorimetry (DSC) analysis was performed on ca. 5 mg samples with a Seiko DSC6200/ EXSTAR6000 apparatus. The temperature readings were calibrated with an indium standard. The DSC measurements were carried out at a scanning rate of 20 °C/min under a nitrogen atmosphere. Usually, the samples were first heated from ambient temperature (25 °C) to 200 °C at a rate of 20 °C/min and subsequently quenched to -140 °C. Then the second scans were run from -140 to 220 °C to record stable thermograms. For CA2.98 and CA2.98-g-PCLs, however, the upper limit of temperature in the heating scans was programmed to be about 260 °C at which the crystallinity of the CA component disappeared swiftly. The cold-crystallization temperature (Tcc), Tm, and Tg were respectively determined from the peak-top position of a crystallization exotherm, that of a melting endotherm, and the midpoint of a discontinuity (baseline shift) in heat flow, in the DSC thermograms obtained in the second heating scan. The change in chemical composition of the specimens hydrolyzed with lipase was determined by measuring 300 MHz 1H NMR spectra. The measuring condition was as follows: apparatus, Varian INOVA 300; solvent, deuterated chloroform (CDCl3), or dimethylsulfoxide (DMSO-d6); solute concentration, 1.0 g/L; internal standard, tetramethylsilane; temperature, 20 °C; pulse width, 3.0 µs; number of scans, 256. Before and after the enzymatic degradation of film specimens, the surface morphology was explored by using a JSPM-5200 atomic force microscope (AFM) in air at 65% RH and 25 °C. An AC mode was utilized with a Nano World PPP-NCHR silicon cantilever of 125 ( 10 µm long and 8 nm tip radius (spring constant, 10-130 N/m). Topography and phase images were obtained simultaneously on an area of 1.0 × 1.0 µm2 at a scan rate of 1 Hz, and typical images were selected after repeating the observation at least three times. High-resolution 13C solid-state NMR measurements were carried out at ambient temperature (20 °C) with a JEOL JNM CMX-300 spectrometer operated at a frequency of 74.7 MHz. The magic angle spinning (MAS) rate was approximately 6 kHz. 13C CP/MAS spectra were obtained in conditions of a contact time 1 ms and a 90° pulse width of 5.0 µs. In quantification of proton spin-lattice relaxation times (T1H), a contact time of 1 ms was used and a proton spin-locking time

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Table 2. Thermal Transition Data for CAs, CBs, PCL, CA-g-PCLs, and CB-g-PCLs Examined in the Present Study samples

Tg (°C)

Tcc (°C)

∆Hcc (J · g-1)

Tm (°C)

∆Hf (J · g-1)

CA2.15a CA2.98a CB2.10 CB2.93a PCL CA2.15-g-PCL2.50 CA2.15-g-PCL9.70 CA2.98-g-PCL2.07

204 175 129 98.3 -61.2 -28.3 -57.2 -54.8 168 -59.0 163 -46.3 -57.4 -38.2 -54.4

n.d.b n.d. n.d. n.d. n.d. n.d. 9.80 n.d. n.d. n.d. n.d. n.d. 9.70 n.d. n.d.

n.d. n.d. n.d. n.d. n.d. n.d. 2.10 n.d. n.d. n.d. n.d. n.d. 5.84 n.d. n.d.

n.d. 295 n.d. n.d. 51.9 n.d. 38.3 n.d. 258 49.5 259 n.d. 48.0 n.d. 51.3

n.d. 18.7 n.d. n.d. 80.7 n.d. 1.55 n.d. 1.57 43.6 0.75 n.d. 43.5 n.d. 42.7

CA2.98-g-PCL9.20a CB2.10-g-PCL2.33 CB2.10-g-PCL9.03 CB2.93-g-PCL3.58 CB2.93-g-PCL12.6a a

Quoted from ref 15.

b

Could not be detected.

τ ranged from 0.1 to 10 s. A total of 1024 scans were accumulated for both 13C CP/MAS spectra and T1H measurements.

Results and Discussion Basic Data of Thermal Transitions. Thermal characterization of the CE-g-PCL products was performed by DSC. Table 2 summarizes transition data for the graft copolymers examined in this study, together with the corresponding data for the original CAs and CBs as trunk polymer and a PCL homopolymer as a reference sample. Any of the CE-g-PCL samples, except for CA2.98-g-PCL copolymers, showed a single Tg varying with oxycaproyl MS between -28 and -58 °C. The two CA2.98-g-PCLs gave another Tg around 165 °C and a small melting endotherm at ∼260 °C, both signals originating from a phase-separated CA domain. This observation is reasonable, because the graft density along the backbone of CA2.98-g-PCL is quite low due to less amount of the residual hydroxyls, and hence, the product virtually behaves like a block copolymer,9,15 giving rise to such a phase separation of the inherently immiscible CA and PCL components. For the block-wise CA2.98-g-PCLs, the phase separation would occur at least in a scale somewhat larger than a couple of tens of nanometers, corresponding to a minimum domain size distinguishable by Tg detection.17,18 In the case of CB2.93g-PCL, contrastively, no segregation behavior was observed from the Tg measurement, because the copolymer (butyryl DS > 2) is composed of an originally miscible pair of trunk and side-chain polymers.15 An additional thermal character found for the CE-g-PCLs of higher MS is a melting endotherm at 38.3-51.3 °C. This result indicates that these PCL-rich compositions allow the PCL side-chains to develop a crystalline phase from the molten state whether the two ingredients are miscible or immiscible; the characterization has already been described in the preceding study.15 Weight Loss Monitoring. Figure 1 compiles weight loss data estimated for film specimens of CE-g-PCLs, CEs, and PCL, each exposed to lipase in phosphate buffer (37 °C, pH ) 7.0). A plain PCL film degraded quickly and showed complete degradation within 24 h. For pristine CAs and CBs, however, no weight loss was detected with exposing time until 120 h. Thus it is found that both acetyl and butyryl groups are hardly degraded by Pseudomonas lipase under the present condition. For the graft copolymers, it was observed that the enzymatic degradation progressed with time to an appreciable extent in

the first 30 h period, but the rate became decreased drastically after an elapse of more than 30 h. Even when 120 h passed, the weight loss of the copolymers was only 3-15% in spite of the occupation of the PCL component at 45-81 wt % to the total polymer matrix (see Table 1). In view of the constraint of the PCL side-chains anchoring onto the cellulosic backbone, such a low weight loss of the CE-g-PCLs may be ascribed to a greater hindrance effect of the CE trunk on the enzymatic attack of lipase to the PCL component. In our previous study on the hydrolysis of CA-g-PLLA with proteinase K,14 it was also shown that the enzymatic attack to the PLLA side-chains was fairly hindered due to the anchoring onto the relatively hydrophobic CA backbone having few unsubstituted hydroxyls. None the less, for the similar hindrance effect of the CA and CB backbones, however, the PCL component should have been degraded, more or less, in the surface region of the film specimens of all the CE-g-PCLs used. As to the CA-based graft series, the weight loss varied seriously with a change in value of either the acetyl DS or oxycaproyl MS. In Figure 1a, we can see that the two CA-gPCLs of MS > 9 exhibit a greater weight loss, compared with the other copolymer of lower MS based on the respective corresponding CA trunks. This result is reasonable; the PCLricher composition would allow the polyester component to form a domain large enough for lipase to access and attack. We can also find an explicit difference regarding the DS dependence, namely, the two CA2.98-g-PCLs showed a weight loss greater than that of the respective CA2.15-g-PCLs of comparable MS. In the former samples, there would be prevalence of a phaseseparated domain structure due to the extremely low graftdensity and the immiscible nature of the CA/PCL pair, as supported by the double Tg data in DSC analysis. As shown in Figure 1b, the CB-g-PCL series showed a somewhat different composition dependence of the weight loss. In comparison between the CB-based grafts of butyryl DS ) 2.10, the weight loss of CB2.10-g-PCL9.03 was larger than that of CB2.10-g-PCL2.33 through the overall exposing time. This situation observed with a common trunk of DS ) 2.10 is similar to that for the CA-based series mentioned above. However, CB2.93-g-PCL3.58 (WPCL ) 52.6 wt %) was digested more rapidly compared with CB2.93-g-PCL12.6 (WPCL ) 79.6 wt %) at least until 30 h odd elapsed. In the latter graft sample of higher MS, a large part of the PCL side-chains crystallize and the rest of this component forms a homogeneous amorphous phase with the CB trunk chains (see Table 2). In contrast, the other film of CB2.93-g-PCL3.58 is a completely amorphous material, where the overall noncrystallizable PCL chains would show a higher degradability without hindrance of enzyme accession by crystalline aggregates. When 60 h and more time passed, however, the weight loss of the PCL-richer CB2.93-g-PCL12.6 film exceeded that of the other one. Comparing the weight loss behavior between the CA-g-PCL and CB-g-PCL series, we readily find that the loss amounts for the CB-based series are generally less than those for the CAbased one. Two factors are responsible for the result. One is the difference in steric hindrance between the acetyl and butyryl substituents. The other is whether the amorphous region in the film sample concerned is miscible or immiscible; this effect should be indicative especially in a comparison using the cellulosic trunks of acyl DS > 2.9. To make such a comparison between CA2.98-g-PCL9.20 and CB2.93-g-PCL12.6, the weight loss value of the former copolymer reached 13.7 wt % in 60 h; whereas only 5 wt % reduction was observed for the latter in the same exposing term. It can be convinced that the steric



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Figure 1. Weight loss for films of (a) CA-g-PCLs and (b) CB-g-PCLs exposed to lipase in phosphate buffer (pH ) 7.0) at 37 °C, plotted as a function of elapsing time. Weight loss profiles of plain PCL, CAs, and CBs as reference samples are also illustrated in both plots.

Figure 2. Monitoring of ∆MS for (a) CA-g-PCLs and (b) CB-g-PCLs exposed to lipase, pursued as a function of elapsing time.

hindrance effect of acyl substituents on the enzymatic attack manifested itself, more strongly in the use of the more bulky CB trunk chain, rather than the use of the CA trunk. In addition, the CB2.93 chains would be mixed, more intimately, with the PCL side-chains due to the miscible character of the CB/PCL pair. In the phase-segregated CA2.98-g-PCL9.20 sample, contrarily, even the inferior hindrance effect of the acetylated cellulosic trunk would be virtually inactive. Compositional Change. To precisely specify the chemical species involved in the weight loss of the CE-g-PCL films exposed to lipase, the compositional change was traced by 1H NMR spectroscopic measurements for the films washed after the prescribed degradation. For CE reference samples, the compositional change was assessed directly through comparison of the acyl DS before and after the hydrolysis. The DS value can be determined from a resonance peak area (A) derived from the methyl protons of acyl groups (1.9-2.3 ppm for CA and 0.8-1.0 ppm for CB) relative to an area (B) of the resonance signals from the protons of glucopyranose unit (2.9-5.2 ppm),6,15 by the following equation

DS ) (A/3)/(B/7)

(1)

By making an actual comparison before and after the 60 h treatment, it was found for all the CEs examined that the acyl DS values reduced by only ∼0.03. This result clearly indicates that the release of acetyl and butyryl groups hardly occurred during the enzymatic degradation experiment. For the graft copolymers, on the other hand, it was difficult to monitor the change of the DS because a C6′′ methylene signal

(4.0-4.2 ppm) from the PCL component overlapped with the resonances of the protons of glucopyranose unit. (Concerning the carbon numbering for the PCL component, see a structural formula shown in Figure 6.) Instead, a resonance signal of C4′′ methylene protons (1.3-1.5 ppm) was clearly separated from that derived from the methyl protons of acetyl or butyryl groups. Therefore, on the basis of the assumption that the acyl groups on the CE backbone are never released during the degradation experiment, we calculated MSn, which is defined as an MS value of a CE-g-PCL film degraded for n h, by the following equation

MSn ) DS × (3C/2A)

(2)

where C denotes a spectral area for the C4′′ methylene protons in PCL side-chains. Then, a change in MS following the enzymatic degradation, ∆MS, was evaluated by the equation

∆MS ) MSn - MS0

(3)

where MS0 is an MS value before the enzymatic hydrolysis started. Figure 2 shows a result of the monitoring of ∆MS for (a) CA-g-PCLs and (b) CB-g-PCLs as a function of exposing time. Evidently, ∆MS values for all the CE-g-PCLs decreased with time in accordance with the weight loss behavior. Just as it was described in the preceding section, the change in MS of CA2.98g-PCL9.20 was of the greatest, ∆MS ) -1.68 being obtained by the hydrolysis continued for 60 h. The weight loss data at the same point was 13.7 wt %, which can be converted to a

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Figure 3. AFM topography images of plain PCL and CE-g-PCL films, observed after enzymatic degradation for 60 h (exceptionally 12 h as to PCL): (a) plain PCL; (b) CA2.15-g-PCL2.50; (c) CA2.98-g-PCL9.20; (d) CB2.10-g-PCL2.33; (e) CB2.93-g-PCL12.6.

∆MS value of -1.60, almost equal to the above data. Similarly, ∆MS values estimated by NMR for the other CE-g-PCLs were in good agreement with the ones calculated from the weight loss data. It is thus ensured for all the CE-g-PCLs that the selective release of PCL segments occurred during the enzymatic hydrolysis, probably in the superficial region of each film specimen. Surface Morphology. AFM observations were carried out to disclose some morphological change in the surface of the

enzymatically degraded films described above. Figure 3 collects AFM topography images of the surfaces for plain PCL and selected CE-g-PCLs; each data was taken after hydrolysis over a period of 60 h except for plain PCL (12 h). With a so-called roughness curve obtained from such a topography image, the root-mean-square roughness (Rrms) was calculated. Table 3 summarizes the Rrms data estimated before and after the enzymatic hydrolysis of each sample. Before starting the enzymatic treatment, all the employed films showed an es-



Table 3. Rrms Data for PCL and CE-g-PCLs, Estimated Before and After 60 h Enzymatic Degradation Rrms (nm) samples

before degradation

after degradation

∆Rrmsa (nm)

PCL CA2.15-g-PCL2.50 CA2.15-g-PCL9.70 CA2.98-g-PCL2.07 CA2.98-g-PCL9.20 CB2.10-g-PCL2.33 CB2.10-g-PCL9.03 CB2.93-g-PCL3.58 CB2.93-g-PCL12.6

3.87 (0.61)b 6.28 (0.74) 4.56 (0.65) 6.17 (0.77) 4.98 (1.24) 5.91 (0.34) 2.49 (0.43) 7.06 (0.61) 3.40 (0.37)

51.2c (1.66) 9.08 (0.71) 7.51 (0.47) 10.3 (0.59) 34.0 (0.79) 6.73 (0.61) 3.40 (0.11) 18.7 (0.94) 15.6 (1.59)

47.3 2.80 2.95 4.13 29.0 0.82 0.91 11.6 12.2

a ∆Rrms ) Rrms(t) - Rrms(0), where Rrms(0) and Rrms(t) represent Rrmss of the test film evaluated before and after degradation for t h, respectively. b Standard deviation of Rrms is given in parentheses. c Evaluated after a 12 h degradation.

sentially smooth surface and Rrms was evaluated as about 3.9 nm for plain PCL and about 2.5-7.1 nm for CE-g-PCLs. As evidenced in Figure 3, all the samples showed, more or less, an undulated surface when degraded for the indicated time, in consequence of the elimination of PCL fragments from the initial film surface. In correspondence with the weight loss data, the surface morphology of the degraded PCL film was considerably rough; the 12 h hydrolysis provided a large Rrms value of 51.2 nm. In the cases of the CE-g-PCLs, the change in Rrms (∆Rrms) following the enzymatic treatment was generally small, reflecting the substantially low degree of degradation of the PCL component linked to the semirigid cellulose backbone; yet we found a comparatively larger ∆Rrms value for graft samples of higher PCL content or with longer PCL side-chains (i.e., higher DPs′) in both of the CA- and CB-based series, as shown in Table 3. Particularly, CA2.98-g-PCL9.20 imparted a ∆Rrms data definitely larger than those for the other copolymer samples. In this kind of standard topography, however, it was difficult to clearly distinguish the difference in surface roughness of the film specimen between the two CE graft series subjected to the PCLselective hydrolysis. Partial abrasion of the surface of polymer films should yield an apparent protuberant structure as relief of the constituent(s) remaining in the superficial region,14 although there would occur

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Table 4. Maximum Position (Hmax) and Standard Deviation (σ) of the Height Distribution of Protuberances for Degraded Films of PCL and CE-g-PCLs samples

Hmax (nm)

σ (nm)

PCL CA2.15-g-PCL2.50 CA2.15-g-PCL9.70 CA2.98-g-PCL2.07 CA2.98-g-PCL9.20 CB2.10-g-PCL2.33 CB2.10-g-PCL9.03 CB2.93-g-PCL3.58 CB2.93-g-PCL12.6

212 4.50 14.6 34.2 170 10.1 12.6 30.8 38.9

33 1.5 4.7 7.1 31 3.4 3.7 7.0 7.4

some reorganization due to aggregations of the same species of molecular chains. The differences in size and regularity of protuberances between the degraded CE-g-PCLs were assessed more quantitatively in terms of a statistical analysis through cross-sectional profiling of the protuberant structure over the whole AFM-scanning surface 1.0 × 1.0 µm2. Figure 4 illustrates histogram plots representing the height distribution of the protuberances observed for the degraded PCL and CE-g-PCLs. In Table 4, standard deviation (σ) values calculated from the distribution data are listed together with the maximum position (Hmax) of the height distribution peaks. In both CA- and CBbased graft series, the distribution peak moves to the side of larger size with an increase in the oxycaployl MS (at a given DS), accompanied by an increase in the deviation σ. This result indicates that the enzymatic degradation of the CE-g-PCLs of higher MS, that is, having longer PCL side-chains, produces deeper abrasions all over the film surface, with a relatively higher extent of irregularity in the magnitude of the protuberant traces. It should also be noted that, when compared between the two block-wise copolymers of WPCL ≈ 80 wt %, CA2.98-gPCL9.20 and CB2.93-g-PCL12.6, a degraded film of the CA-based product showed a more eroded surface (Hmax ≈ 170 nm) relative to that (Hmax ≈ 39 nm) of the corresponding film of the CBbased one. This observation supports that the former film sample is capable of forming a considerably larger PCL domain, differing from the latter composed of a miscible constituent pair. Subsequently, we provided further insight into the surface morphology of CE-g-PCL films by using AFM phase images

Figure 4. Histograms showing height distribution of the protuberances formed on the individual surfaces of degraded films of (a) CA-g-PCLs and (b) CB-g-PCLs.

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Figure 5. AFM phase images of plain PCL and CE-g-PCL films: (a) plain PCL; (b) CA2.15-g-PCL2.50; (c) CA2.98-g-PCL9.20; (d) CB2.10-g-PCL2.33; (e) CB2.93-g-PCL12.6. The left side (a-e) and right side (a′-e′) are the phase images, observed respectively before and after enzymatic degradation for 60 h (or 12 h for plain PCL). Table 5. T1H Values for CA2.98-g-PCL9.20 and CB2.93-g-PCL12.6 Samples T1H (s) CA or CB component

PCL component

samples

C2/C3/C5

C2′

C4′

C2′′

C5′′

CA2.98-g-PCL9.20 CB2.93-g-PCL12.6

0.80 0.65

0.82 n.d.a

0.67

0.90 0.67

0.90 0.68

a

Could not be detected.

before and after the hydrolysis. Generally, this kind of image is known to reflect differences in the chemical and mechanical properties such as adhesiveness, viscoelasticity, friction, and so on. In the present case, the phase image may be regarded as a high-resolution map characterizing the distribution of CE and PCL domains in the superficial region of each film sample. Figure 5 exemplifies the actual data for plain PCL, and CAand CB-g-PCLs; the respective five pairs of micrographs were obtained before and after the degradation of the film specimens for 12 h (PCL) or 60 h (others). In Figure 5a for the undegraded PCL film, we can see a contrast of dark and bright phases arranged in an irregularly banded fashion. Considering the intolerable habit of crystal development of this aliphatic polyester, it is reasonably inferred that the bright areas correspond to the rigid crystalline phase. After enzymatic degradation of the film for 12 h, lamella-like structures became pronounced, as shown in Figure 5a′. It can

be taken that the preferential degradation of the amorphous regions19 threw the PCL lamellar crystals into relief on the film surface. The phase images of CE-g-PCL films also varied drastically between before and after the degradation. For undegraded CA2.15-g-PCL2.50 (Figure 5b) and CB2.10-g-PCL2.33 (Figure 5d), a granular texture was observed, in which bright and dark phases were mingled with each other so finely that the boundary was obscured. Taking into account their compositions of lower MS and therefore no melting signal in the DSC thermograms (see Table 2), the bright and dark areas may be tentatively assigned to a more rigid CE domain and a flexible PCL one, respectively. After the 60 h degradation process, it was found for both copolymers that the profile of the bright areas became sharpened, as shown in Figure 5b′,d′. Through comparison between the phase and the topography images after the degradation, it was confirmed that the bright areas in the phase images corresponded substantially to the protuberances as remains in the topography ones. Therefore, it can be rationalized that the bright phase enriched after the degradation is identified as the CE domain bared on the film surface by the selective enzymatic hydrolysis of the PCL component. Then the domain size was estimated as 8-12 nm for CA2.15-g-PCL2.50 and 10-15 nm for CB2.10-gPCL2.33. Here, it should be remarked that such a domain size evaluated from the phase image does not necessarily coincide with the apparent size of the protuberant structure observed in



2837

Figure 6. Solid-state 13C CP/MAS NMR spectra for CA2.98-g-PCL9.20 and CB2.93-g-PCL12.6 and their peak assignments. Asterisks denote a spinning sideband overlapping with a resonance signal of C4 pyranose carbon. The C2′′ peak of CB2.93-g-PCL12.6 contains a negligible intensity of C2′ of the butyryl substituent.

the topography image; the surface roughness of polymer-eroded films is solely emphasized in the latter mode. The phase images of undegraded films of CA2.98-g-PCL9.20 and CB2.93-g-PCL12.6 are displayed in Figure 5c and e, respectively. Considering that the PCL component in these copolymers can crystallize owing to the PCL-rich compositions (WPCL ≈ 80 wt %), the brighter areas may be ascribed mainly to the PCL crystal domains. In this assumption, we estimated from the images that the CB2.93-g-PCL12.6 sample developed a lamellalike morphology of 40-60 nm in width and 140-280 nm in length, whereas no such structure was formed in the CA2.98-gPCL9.20 sample. This estimation is consistent with the previous observation15 that CB2.93-g-PCLs rich in PCL exhibited larger spherulites having periodically banded extinction rings under a polarized optical microscope, while CA2.98-g-PCL9.20 gave a different texture composed of numerous minute grains (fine spherulites). After enzymatic treatment, as can be seen from Figure 5c′, the degraded CA2.98-g-PCL9.20 showed an assembly morphology of considerably large domains of 25-45 nm in diameter size, deservedly belonging to the remaining CA component. This result strongly suggests that the trunk and side-chain polymers of CA2.98-g-PCL9.20 were initially phase-separated at least on the scale of a couple of tens of nanometers, even though the degradation process might have promoted the phase segregation due to some cohering rearrangement of the CA component as residue. The marked domain formability originating from the extremely low graft-density and the intrinsically poor miscibility of the CA/PCL pair should lower the hindrance effect of the CA constituent on the enzymatic attack onto the PCL chains, which results in the higher degradability of the PCL component (Figure 1a). On the other hand, for the degraded CB2.93-gPCL12.6, the domain size of the CB component was estimated roughly as 10-20 nm. As shown in Figure 5e′, however, the boundaries between the individual domains were much more indistinctive than in the case of the CA component of CA2.98g-PCL9.20. This observation satisfies us that the PCL degradability of the CB2.93-based sample was rather low (see Figure 1b) due to the higher hindrance effect of the trunk component miscible with PCL, even in such a composition with an extremely low graft-density and longer PCL side-chains. Quantification of Domain Size by T1H Measurement. As is well established, 1H spin-lattice relaxation time (T1H) measurements in solid-state 13C NMR spectroscopy provide

information about some microphase structure of polymers in a scale of a few tens of nanometers. The T1H measurements for specific carbons in a graft copolymer will also make it possible to identify the microheterogeneous structure through detection of the difference in the relaxation behavior between the trunk and side-chain polymer constituents. In the present work, we carried out the measurements for virgin films of CA2.98-g-PCL9.20 and CB2.93-g-PCL12.6, which formed a striking contrast to each other in the AFM phase image both before and after the surface degradation and estimated the heterogeneity of the constituents in a scale of length of 1H spin-diffusion. T1H values can be obtained practically by fitting the carbon resonance intensity to the following single-exponential equation

M(τ) ) M0{1 - 2 exp(-τ/T1H)}

(4)

where M(τ) is the magnetization intensity observed as a function of the spin-locking time τ and the initial intensity is given by M(0) ) -M0. As a general rule, if two polymer components are in a homogeneous mixing state on the scale over which 1H spin diffusion can take place in a time T1H, the T1H values for different protons belonging to the respective components may be equalized to each other. The result of the T1H measurements for the two CE-g-PCLs of acyl DS > 2.9 is summarized in Table 5. 13C CP/MAS spectra of the two graft copolymers and the peak assignments for the constituents are also shown in Figure 6. The monitoring of the relaxation process was conducted for the peak intensities of C2/C3/C5 pyranose carbons and C2′ (or C4′) methyl carbons of the CA (or CB) component and for those of C2′′ and C5′′ methylene carbons of the PCL component. As for CA2.98-g-PCL9.20, we obtained T1H ≈ 0.81 s for the CA component and a comparatively larger value 0.90 s for the PCL component. This implies that the sizes of the CA and PCL domains are comparable to, or rather larger than the maximum diffusive path length L that is given by the equation20

L = (6DT1H)1/2

(5)

where D is the diffusion coefficient, usually taken to be ∼10-12 cm2/s in organic polymer materials. Simple application of this relation with the T1H data of 0.81 and 0.90 s leads to an estimation of L ≈ 22.0 and 23.2 nm, respectively. Therefore,

2838


conversely, we can infer that the two components constituting the CA2.98-g-PCL9.20 sample would be phase-separated on the scale of about 23 nm, before exposure to lipase in phosphate buffer solution. The phase-separated scale of >23 nm is consistent with the apparent CA domain size of 25-45 nm roughly estimated from the AFM phase image (Figure 5c′) for the surface-degraded film of the same copolymer. Thus, it follows that the CA domain, originally invisible to the AFM due to the PCL-rich composition, was successfully visualized by the selective release of the PCL component in the superficial region of the film. On the other hand, as to the CB2.93-g-PCL12.6 sample, we obtained noticeably shorter values of T1H: 0.65-0.67 s for the CB component and 0.67-0.68 s for the PCL component. Obviously, the two data are very close to each other, indicating that the two polymer constituents are coexistent in a range where the mutual spin diffusion is permitted over a period of the homogenized T1H. By substituting an averaged T1H of 0.67 s into eq 5, it is found that the CB and PCL components are intimately mixed on a scale of less than 20 nm. This result also supports that the enzymatic attack onto the PCL side-chains in the graft copolymer would be so hindered by the hydrophobic CB trunk that is originally miscible with PCL.

Conclusions Enzymatic hydrolysis was conducted for different compositions of CA-g-PCL and CB-g-PCL by employing Pseudomonas lipase. The selective hydrolysis behavior was characterized by monitoring the weight loss, compositional change, and surface morphology of film specimens of the graft copolymers, while they were exposed to the enzyme. A possible heterogeneity in some of the copolymer films, which was signaled by the degradation, was also inspected by T1H measurements in solidstate 13C NMR spectroscopy. For both copolymer series, a hindrance effect of the CE trunks on the enzymatic attack to the PCL component was observed; exceptionally, however, the effect was rather inactive in blocklike CA-g-PCLs of acetyl DS ) 2.98 due to a noticeable domain formability that can be ascribed to the extremely low graftdensity as well as to the intrinsically poor miscibility of the CA/PCL polymer pair. After the selective hydrolytic degradation of the PCL component, the AFM topography revealed that the CA and CB constituents as residues formed a protuberant structure having an average altitude of ca. 5-170 nm, variant depending on the copolymer architecture, on the surface of the respective film specimens. A hydrolyzed film of blockwise

Kusumi et al.

CA2.98-g-PCL9.20 showed particularly larger CA domains exceeding a 25 nm size in the phase image, which was consistent with the estimation of a heterogeneity scale of >23 nm by T1H measurements for the initial intercomponent mixing state. Looking through the accumulated data of degradation behavior, we can reasonably conclude that the present PCL-grafted cellulosic copolymers may be categorized as a “spatiotemporally biodegradation-controllable” material, because not only the degradation rate but also the surface morphology of the molded films was proved to be subtly changeable by adequate selections of the compositional parameters and intercomponent miscibility of the original products. The result of this study should be useful for expanding the availability of cellulosics as a major component to design novel polymeric materials furnished with highfunctionality.

References and Notes (1) Klemm, D.; Heublein, B.; Fink, H. P.; Bohn, A. Angew. Chem., Int. Ed. 2005, 44, 3358–3393. (2) Nishio, Y. AdV. Polym. Sci. 2006, 205, 97–151. (3) Nishio, Y.; Manley, R. S. J. Macromolecules 1988, 21, 1270–1277. (4) Buchanan, C. M.; Gedon, S. C.; White, A. W.; Wood, M. D. Macromolecules 1992, 25, 7373–7381. (5) Nishio, Y.; Matsuda, K.; Miyashita, Y.; Kimura, N.; Suzuki, H. Cellulose 1997, 4, 131–145. (6) Kusumi, R.; Inoue, Y.; Shirakawa, M.; Miyashita, Y.; Nishio, Y. Cellulose 2008, 15, 1–16. (7) Ohno, T.; Nishio, Y. Macromolecules 2007, 40, 3468–3476. (8) Teramoto, Y.; Nishio, Y. Polymer 2003, 44, 2701–2709. (9) Teramoto, Y.; Ama, S.; Higeshiro, T.; Nishio, Y. Macromol. Chem. Phys. 2004, 205, 1904–1915. (10) Vide´ki, B.; Kle´bert, S.; Puka´nszky, B. J. Polym. Sci., Part B: Polym. Phys. 2007, 45, 873–883. (11) Dong, H.; Xu, Q.; Li, Y.; Mo, S.; Cai, S.; Liu, L. Colloids Surf., B 2008, 66, 26–33. (12) Edgar, K. J.; Buchanan, C. M.; Debenham, J. S.; Rundquist, P. A.; Seiler, B. D.; Shelton, M. C.; Tindall, D. Prog. Polym. Sci. 2001, 26, 1605–1688. (13) Teramoto, Y.; Nishio, Y. Biomacromolecules 2004, 5, 397–406. (14) Teramoto, Y.; Nishio, Y. Biomacromolecules 2004, 5, 407–414. (15) Kusumi, R.; Teramoto, Y.; Nishio, Y. Macromol. Chem. Phys. 2008, 209, 2135–2146. (16) Gan, Z.; Liang, Q.; Zhang, J.; Jing, X. Polym. Degrad. Stab. 1997, 56, 209–213. (17) Utracki, L. A. Polymer Alloys and Blends; Hanser: Munich, 1990. (18) Nishio, Y. In Cellulosic Polymers, Blends, and Composites; Gilbert, R. D., Ed.; Hanser: Munich, 1994; Chapter 5, pp 95-113. (19) Mochizuki, M.; Hirano, M.; Kanmuri, Y.; Kudo, K.; Tokiwa, Y. J. Appl. Polym. Sci. 1995, 55, 289–296. (20) McBrierty, V. J.; Douglass, D. C. J. Polym. Sci., Macromol. ReV. 1981, 16, 295–366.

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