Comparative study on the enzymatic degradation of poly(lactic-co-glycolic acid) by hydrolytic enzymes based on the colorimetric quantification of glycolic acid

Polymer Testing 30 (2011) 743–748

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Analysis Method

Comparative study on the enzymatic degradation of poly (lactic-co-glycolic acid) by hydrolytic enzymes based on the colorimetric quantification of glycolic acid Michael Kemme*, Ines Prokesch, Regina Heinzel-Wieland Department of Chemical Engineering and Biotechnology, Hochschule Darmstadt, University of Applied Sciences, Schnittspahnstrasse 12, D-64287 Darmstadt, Germany

a r t i c l e i n f o

a b s t r a c t

Article history: Received 4 May 2011 Accepted 24 June 2011

The enzyme-catalysed cleavage of ester bonds of poly(lactic-co-glycolic acid) (PLGA) has been monitored by an improved approach for the quantification of glycolic acid as indicator of copolymer hydrolysis. Glycolic acid released into the degradation medium was determined colorimetrically using chromotropic acid. The assay established is sensitive, as well as being both rapid and economical. The proposed method was applied to assess enzymatic degradation of a series of PLGA polyesters containing different proportions of monomers, end groups of PLGA chains and molecular weights. Among 22 commercially available hydrolytic enzymes (i.e. esterases, lipases and proteases) from different sources, the lipases from Candida antarctica, Candida cylindracea, Candida rugosa, Mucor miehei, Rhizopus arrhizus and porcine pancreas and the esterase from M. miehei all had a significant effect on PLGA degradation, often increasing the rate of ester bond cleavage by a factor of 25 compared to non-enzymatic hydrolysis. The most remarkable substrate specificity was observed for C. antarctica lipase with the rate of degradation being directly proportional to the glycolic acid content. In contrast, degradation by M. miehei esterase and R. arrhizus lipase were nearly independent of PLGA structure. Ó 2011 Elsevier Ltd. All rights reserved.

Keywords: Chromotropic acid Enzymatic degradation Esterase Glycolic acid Lipase Poly(lactic-co-glycolic acid)

1. Introduction Within the frame of sustainable development, aliphatic polyesters based on lactic or glycolic acid and their copolymers have been extensively investigated due to their biocompatibility and biodegradability [1]. In particular, poly(lactic-co-glycolic acid) (PLGA) has received considerable attention for use in temporary biomedical applications (e.g. drug-delivery systems, bioabsorbable sutures, bone fixation devices and tissue engineering scaffolds), but also as an industrial product for environmentally friendly packaging systems and single-use disposable items [2]. It is

* Corresponding author. Tel.: þ49 (0) 6151 16 8633; fax: þ49 (0) 6151 16 8404. E-mail address: [email protected] (M. Kemme). 0142-9418/$ – see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.polymertesting.2011.06.009

generally accepted that PLGA demonstrates desirable mechanical properties, good toxicological safety and tuneable degradation rates which are affected by several physical and chemical parameters, such as initial pH, ionic strength and temperature of external medium, the molar ratio of monomers, enantiomeric form, initial molecular weight, crystallinity and specimen size [3,4]. The abiotic degradation of a solid PLGA matrix in an aqueous environment can occur through bulk erosion involving hydration of amorphous polymer regions, passive ester bond hydrolysis, significant mass loss and solubilisation of oligomers with release of lactic and glycolic acid, both incorporated into cellular intermediate metabolism and assimilated in vivo [5]. As far as biodegradability is concerned, it was reported that PLGA fragmentation could be significantly accelerated by different microbial species which assimilated the

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degradation products as sole carbon source [6]. This would suggest the involvement of either intracellular enzymes or enzymes secreted from bacteria, yeast or fungi [7]. The evaluation of the microbial degradation process and mechanism requires fundamental understanding of the enzymatic activities which take place through adsorption of the enzyme on the surface of the polymer substrate followed up by hydrolysis of the polyester main chain. In contrast to reports on poly(lactic acid) (PLA), little is known about specific enzymes (i.e. esterases, lipases, proteases) from microbial origin responsible for cleavage of ester bonds between lactic and glycolic acid monomers in the PLGA backbone. Enzymatic hydrolysis of the copolymer has been investigated by few researchers, especially in the presence of the mammalian protease trypsin and proteinase K, a proteolytic enzyme secreted from the fungus Tritirachium album [8,9]. Additionally, two kinds of fungal lipases were reported to promote degradation of PLGA, i.e. Rhizopus arrhizus lipase [9,10], and Rhizopus delemer lipase [11]. However, there is a clear difficulty in comparing and explaining the degradation rates of the enzymes used in these studies due to the lack of standardization of reaction conditions and assay systems. Studies on degradation of raw PLGA samples have employed a broad range of techniques including gravimetry, viscosimetry, total organic carbon analysis of watersoluble fraction, size-exclusion chromatography, liquid chromatography of hydrolysis products, differential scanning calorimetry, scanning electron microscopy and X-ray diffraction [12]. These procedures, however, either are not readily applicable to small-scale studies or lack the accuracy and specificity required for assessing subtle variations in hydrolytic reactivity towards PLGA copolymers with different monomer compositions. More precise and rapid colorimetric methods have been described for the quantitative analysis of lactic acid released in aqueous medium [13]. The main limitation of the latter technique is that only the L-monomer is determined in samples with D,L-stereoisomers of lactic acid. Furthermore, since several factors of the enzyme-catalysed PLGA hydrolysis, such as temperature and enzyme concentration, play an important role in the polymer degradation rate, it is important to establish a reproducible and easy-handling method free of steric constraints suitable for processing large series of samples from multiple point degradation tests. In this paper, we report on the enzymatic biodegradation of PLGA copolymers with different amounts of glycolic acid using 22 commercially available hydrolytic enzymes. The aim of this work was to identify enzymes, especially from microbial origin, that are capable of rapid and complete ester-linkage scission. In order to accurately monitor the enzymatic degradation of PLGA and to overcome the analytical limitations of classical techniques, an improved colorimetric method based on the quantitative assay of glycolic acid released in the incubation medium is reported. The low-cost method established is simple with high precision, which is promising for testing of PLGA biodegradation in research and industry.

2. Experimental 2.1. Materials The PLGA copolymers used in this study were commercial RESOMERÒ products made by Boehringer Ingelheim (Table 1). Except for RG 503 (a kind gift of Roland Klein, Darmstadt), the other members of the PLGA series were purchased from Sigma–Aldrich (Steinheim, Germany) and were applied as received. Glycolic acid, p-nitrophenylacetate (pNPA) and p-nitrophenyl-palmitate (pNPP) were also from Sigma–Aldrich. Chromotropic acid was obtained from Roth (Karlsruhe, Germany) and the Pierce protein assay kit using bicinchoninic acid (BCA) was from Thermo Scientific (Rockford, IL, USA). All other solvents and chemicals, unless specified, were commercial products of the highest grade available. A number of different commercially available hydrolytic enzymes (Table 2) were tested with respect to their degradation potential against PLGA. The supplied preparations were used in the experiments without further purification. Stock solutions of lipases were prepared by dissolving the enzyme formulations in 0.1 M sodium phosphate, pH 7, at a concentration of 5–20 mg/ml (solid mass). Each esterase was dissolved in 50 mM Tris–HCl, pH 8.0, to a concentration of 4–10 mg/ml. Proteases were dissolved to a concentration of 5–10 mg/ml in water or 1 mM HCl for trypsin and chymotrypsin, respectively. All enzyme solutions were stored in aliquots at 25  C. Prior to use, each enzyme formulation was assayed for its protein content and for its esterolytic and lipolytic activity. 2.2. Enzymatic degradation Enzymatic degradation studies were conducted in 50 mM Tris–HCl, pH 7.4, buffer containing 150 mM NaCl and 0.02% (w/v) Tween 20 as release medium. Samples of powdered polymer (5 mg each) were dispersed in 2-ml reaction tubes in 1 ml of release medium. After addition of 0.1 ml enzyme solution (1 mg/ml) in release medium, the reaction mixtures were incubated on a magnetic stirrer (800 rpm) at constant temperature of 37  C. To observe concentration and temperature dependent changes in PLGA degradation, samples with 25, 50 and 100 mg enzyme were thermostated at 28, 37 and 50  C. At predetermined time intervals, the tubes were centrifuged at 10,000 x g (Heraeus Biofuge pico, Kendro, Langenselbold, Germany) for 10 min, 1 ml of the supernatants removed and stored at 25  C until glycolic acid determination. The remaining pellets were redispersed in 1 ml fresh buffer/enzyme media to restore the original level of enzymatic activity.

Table 1 PLGA ResomerÒ copolymers investigated comparatively. PLGA type

D,L-lactic:glycolic acid ratio (molar)

Molecular weight range

End group

RG RG RG RG

50:50 50:50 65:35 75:25

24,000–38,000 24,000–38,000 24,000–38,000 4000–15,000

alkyl ester free carboxylic acid free carboxylic acid free carboxylic acid

503 503 H 653 H 752 H

M. Kemme et al. / Polymer Testing 30 (2011) 743–748

Enzyme (abbreviation)

Origin

Bacillus species esterase (BsE) Mucor miehei esterase (MmE) Porcine liver esterase (PLE) Aspergillus niger lipase (AnL) Aspergillus species lipase (AsL) Candida antarctica lipase (CaL) Candida cylindracea lipase (CcL) Candida rugosa lipase (CrL) M. miehei lipase (MmL) Pseudomonas cepacia lipase (PcL) Pseudomonas fluorescens lipase (PfL) Porcine pancreatic lipase (PPL) Rhizopus arrhizus lipase (RaL) Rhizopus niveus lipase (RnL) a-Chymotrypsin, bovine pancreasa Collagenase, Clostridium histolyticumb Pronase, Streptomyces griseus Proteinase K, Tritirachium album Proteinase N, Bacillus subtilis Proteinase VIII, Bacillus licheniformis Subtilisin, Bacillus licheniformis Trypsin, bovine pancreas

bacteria fungus mammalian fungus fungus yeast yeast yeast fungus bacteria bacteria

Protein Activity content (U/mg) (%) pNPA pNPP 0.3 6.5 100.0 0.5 0.7 0.7 8.1 45.9 1.6 10.7 10.5

< 0.1 3.9 40.0 < 0.1 < 0.1 7.9 9.1 9.7 0.8 353.0 256.0

mammalian 39 fungus 28 fungus 38 mammalian 100

0.2 3.3 0.1 –c

0 0 0 nd

bacteria

55

< 0.1 nd

bacteria fungus bacteria bacteria

67 100 50 100

33 8 50 34 24 35 15 20 55 27 15

bacteria 100 mammalian 100

< 0.1 1.2 < 0.1 1.2

nd 0 nd nd

11.6 nd < 0.1 nd

nd, not determined. The hydrolytic enzymes were delivered from Sigma-Aldrich. a Delivered from Merck (Darmstadt, Germany). b Deliverd from Serva (Heidelberg, Germany). c The sample was found to contain 75% active enzyme as determined by active-site titration with pNPA.

Controls were realised for each polymer in release medium without enzyme. 2.3. Colorimetric measurements Glycolic acid released during the degradation of PLGA was determined by a modified colorimetric method [13,14]. In summary, 100 ml sample solution was mixed with 2 ml of freshly prepared 0.1% (w/v) chromotropic acid in concentrated sulphuric acid and incubated at 95  C for 30 min. After cooling on ice for 5 min, the reaction was terminated by adding 4 ml ice-cold water and absorbance read at 578 nm using a Ultrospec 2100 pro spectrophotometer (Amersham Pharmacia Biotech, Uppsala, Sweden). The linearity of the method was verified by a calibration graph based on pure glycolic acid standards from 0 to 0.5 mg/ml. The observed glycolic acid amounts were corrected by subtraction of the respective control values for nonenzymatic PLGA hydrolysis. The biodegradability (%) of copolymers was calculated by dividing cumulative release rates of glycolic acid in all withdrawn samples with total glycolic acid amount calculated from polymer composition. Lipolytic activity was assayed spectrophotometrically in aqueous solution according to a previously published method [15] with some modifications using pNPP as the substrate. The enzyme solution (100 ml) was emulsified for 10 min at 37  C in 800 ml of phosphate buffer (50 mM, pH 8.0) containing 4% (w/v) Triton X-100 and 0.2% (w/v) gum

arabic. The reaction was started by adding 100 ml 10 mM pNPP in isopropanol and the absorbance measured at 410 nm for 5 min. One unit (U) of lipase activity was defined as the amount of enzyme necessary to release 1 mmol p-nitrophenol per minute. Esterolytic activity was measured by hydrolysis of pNPA in a routine assay [16]. Protein concentration was determined by the BCA protein assay kit using the standard protocol with bovine serum albumin as a calibration reference. 3. Results and discussion Biomedical applications and the huge potential for biodegradable packaging systems based on PLGA copolymers require fundamental and comprehensive information concerning their enzymatic hydrolysis because interactions between polyester and enzyme are inevitable during exposure to body fluids, cells and microorganisms in natural environments. A particular problem arises in monitoring the degradation of PLGA in small-scale hydrolysis studies, where conventional techniques are impractical [12]. In order to accurately define the enzyme-accelerated hydrolysis of PLGA, a rapid and simple colorimetric method was established which allowed the sensitive quantitation of glycolic acid as degradation end product. 3.1. Analytical validation A commonly used colorimetric method for determination of glycolic acid in urine [14] was adapted to small-scale samples of PLGA hydrolysis. The assay protocol was based on the oxidation of glycolic acid to formaldehyde in concentrated sulphuric acid and the subsequent colour reaction between formaldehyde and chromotropic acid yielding red purple condensation products. Lactic acid is known not to interfere with this test [14]. The linearity of the method was evaluated by analysing a series of glycolic acid standards (Fig. 1). The results were subjected to best-fit linear regression giving y ¼ 0.0033x þ 0.0062 as the equation for calculating glycolic acid (x) released. The coefficient of determination (R2) of 0.9991 indicates a very strong correlation between analytical data [17] as observed in the present investigation. The standard calibration curve of the assay (Fig. 1) demonstrated highly linear results over glycolic acid concentrations in the range of 25–500 mg/ml. The Absorbance (578 nm)

Table 2 Characteristic data of enzymes used.

745

1.5

1

0.5

y = 0.0033x + 0.0062 R2 = 0.9991

0 0

100

200

300

400

500

600

Glycolic acid concentration (µg/ml) Fig. 1. Calibration curve for the quantitative determination of glycolic acid by the colorimetric assay with chromotropic acid based on eight replicate experiments.

M. Kemme et al. / Polymer Testing 30 (2011) 743–748

The role of enzymes in any PLGA biodegradation is unclear. Most of the literature indicates that the scission of ester bonds between lactic and glycolic acid does not involve enzymatic activity and is solely through hydrolysis. However, some reports have suggested an enzymatic role in PLGA breakdown based on the differences between in vitro and in vivo degradation rates [18]. To gain preliminary information about the degradation behaviour of PLGA under different conditions in the presence of enzyme, the lipase from Candida antarctica (CaL, Table 2) was chosen for extended systematic investigations. Glycolic acid production as an indicator of polymer degradation was detected based on the colorimetric assay. As a first step, different CaL concentrations were tested in ResomerÒ RG 503 suspensions at 37  C in buffered saline at pH 7.4 (Fig. 2A). A control without enzyme was run as reference indicating no significant non-enzymatic degradation. In contrast, the hydrolysis of PLGA took place in all cases in the presence of CaL, demonstrating a time course with a hyperbolic release profile of monomer. The degradation rate was very high at the beginning and decreased with time, becoming almost constant after 24 h. The reaction was nearly complete within 48 h.Increasing the amount of enzyme from 25 to 100 mg in 1.1-ml reaction volume resulted in increased liberation of glycolic acid. Therefore, in the subsequent PLGA degradation experiments, all reactions were carried out with 100 mg enzyme/ ml incubation mixture. The influence of temperature on the enzymatic degradation of ResomerÒ RG 503 by CaL is presented in Fig. 2B. Lipase degraded PLGA in a temperature-dependent manner, with the rate of degradation being inversely proportional to the temperature. The maximum degradation is observed at 28  C, while at higher temperatures, 37 and 50  C, a large decrease in the enzymatic hydrolysis is seen. Even though, CaL showed maximal activity at 35  C towards small lipid substrates in the presence of a high-molecular-weight PLGA substrate, with similar activities between 20 and 40  C [19], lower temperatures favoured scission of polyester bonds. This effect can be explained based on the molecular motion of the copolymer chains. At 28  C, perhaps the molecular mobility of the polyester chains is such that the maximum amount of lipase might have reached the polymer surface and resulted in the highest degradation, whereas beyond this temperature mobility of the polyester chains has increased and might have hindered enzyme molecules reaching the ester bonds. Taken together, temperature and concentration dependence of CaL-mediated glycolic acid release provide a strong argument for an enzymatic role in PLGA biodegradation. The effect of the type of end groups on the enzymatic degradation of PLGA was investigated by selecting ResomerÒ RG 503 and RG 503 H (Fig. 2C). Both co-

Biodegradability (%)

3.2. Evaluation of PLGA biodegradation

A 100

Control 25 µg/ml 50 µg/ml 100 µg/ml

80 60 40 20 0 0

B Biodegradability (%)

pure glycolic acid solutions and samples from PLGA degradation mixtures showed unchanged colours. The closeness of these results underlined that the proposed method represents an appropriate tool for assaying PLGA hydrolysis quantitatively.

8

16 24 32 Degradation time (h)

40

48

100 28 °C 37 °C 50 °C

80 60 40 20 0

0

8

16

24

32

40

48

Degradation time (h)

C Biodegradability (%)

746

100 RG 503 RG 503 H RG 653 H RG 752 H

80 60 40 20 0 0

8

16 24 32 Degradation time (h)

40

48

Fig. 2. Time course of CaL-mediated PLGA degradation in ResomerÒ suspensions at pH 7.4. Degradation profiles of RG 503 with different enzyme concentrations at 37  C (A) and at different temperatures with 100 mg/ml CaL (B). Time-dependent biodegradability of ResomerÒ samples with 100 mg/ml CaL at 37  C (C).

polyesters with equimolar monomer composition contain alkyl esters and free carboxylic acid groups as chain terminators (Table 1). The degradation rates of PLGA with free acid ends were reduced by almost half compared to the copolymer with blocked end groups. The possible reason for the slow hydrolysis of ResomerÒ RG 503 H might be the electrostatic arrangement of the negatively charged acid groups which seems to affect the enzyme attack adversely. In order to investigate the influence of the co-monomer composition on PLGA hydrolysis by CaL, the degradation of the three ResomerÒ types RG 503 H, RG 653 H and RG 752 H

M. Kemme et al. / Polymer Testing 30 (2011) 743–748

with identical end groups but different molar ratios of lactic and glycolic acid (50:50, 65:35 and 75:25, respectively; see Table 1 for the properties of polymers) was compared (Fig. 2C). All the PLGA showed significant release of glycolic acid with degradation time. The ResomerÒ RG 503 H degraded faster than RG 653 H, whereas RG 752 H showed less degradation compared to the other two polymers. This phenomenon was due to the higher content of hydrophilic glycolic acid residues in RG 503 H, which could facilitate the water absorption and diffusion necessary for the lipasecatalyzed cleavage of ester bonds. This was consistent with literature data on non-enzymatic PLGA degradation, indicating enhanced susceptibility of the chain scission reaction due to the increase in glycolic acid content within the copolymer [20]. Additionally, we found that the release of glycolic acid from RG 752 H and RG 653 H was similar in both cases, although the molecular-weight range of the PLGA types differed significantly (Table 1). The result suggests that there is no obvious relationship between the chain length of PLGA and the amount of degradation. 3.3. Comparison of hydrolytic enzymes Knowledge of the biodegradation behaviour of PLGA under different conditions is really important in order to understand the co-polyester breakdown in natural environments. Due to water insolubility and the size of the polymer molecules, microorganisms are not able to pick up PLGA directly into the cells where most of the metabolic pathways take place, but first have to secrete extracellular enzyme(s) which depolymerize the co-polyester outside the cells. The resulting water soluble intermediates can than be taken into the microorganism. The reduction of chain length based on enzyme triggered mechanisms requires the assistance of members of the hydrolase family consisting, amongst others, of esterases, lipases and proteases. A systematic study of the enzymatic degradation of PLGA has not yet been reported in the literature. For the first time, we have comprehensively investigated the substrate specificity of three esterases, eleven lipases, and eight proteases, mainly from microbial origin (Table 2), on ResomerÒ types with different molar ratios of lactic and glycolic acid in buffered suspensions at physiological pH. The extent of PLGA degradation was determined by quantifying the released glycolic acid residues with the presented colorimetric assay. All experiments were carried out for 48 h, enzyme concentration being maintained at 100 mg/ml. Prior to biocatalytic PLGA cleavage, the activity of all enzyme formulations was verified by esterolytic and lipolytic assays (Table 2). Out of the tested enzymes, lipases constitute an important group of the hydrolase family for enzymatic degradation of aliphatic polyesters by cleaving ester bonds randomly along the main chain of the polymer substrates [21]. Significant degradation of PLGA was seen only in samples digested with CaL, Candida cylindracea lipase (CcL), Candida rugosa lipase (CrL), porcine pancreatic lipase (PPL), and R. arrhizus lipase (RaL), while the lipase from Mucor miehei (MmL) degraded all PLGA samples only at a low rate (Fig. 3).

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CaL exhibited the highest activity under the test conditions by cleaving almost 95% of ResomerÒ RG 503, increasing the rate of ester bond hydrolysis by a factor of 25 compared to the non-enzymatic reaction. Additionally, the most remarkable substrate specificity was observed for CaL with the rate of degradation being directly proportional to the glycolic acid content of PLGA. For the degradation of different ResomerÒ types by lipases, the activity orders were as follows: CaL > CcLwRaL > CrL > PPL > MmL (RG 503), CaL > RaL > PPL w CrL > CcL > MmL (RG 503 H), RaL w CaL > CcL > CrLwPPL > MmL (RG 653 H), RaL > CaL > PPL > CrL > MmL > CcL (RG 752 H). While the varying activities of the tested lipases preclude a comparative and quantitative analysis of the effect of these enzymes, it was clear that microbial lipases are able to influence PLGA degradation under some conditions. The difference in the catalytic performance of the lipases towards PLGA could be explained by different binding capacities of the enzymes, since it is known that each lipase has the same catalytic behaviour but the binding to substrates with structural variations around the ester linkage varies depending on the microbial source of enzyme [22]. In polymer suspensions, lipases are acting on an unusual solid substrate, which is different to their normal mode of action at oil/water interfaces to hydrolyse primarily triacylglycerols. Nevertheless, significant amounts of hydrolysis were observed for CaL, CcL, CrL, PPL, and RaL promoting efficient degradation of PLGA. Among three esterases, only M. miehei esterase (MmE) showed remarkable degradability of the four ResomerÒ types (Fig. 3). It seemed that the rate of ester bond hydrolysis exhibited similar trends independently of the composition of PLGA. As esterases catalyse the cleavage of water soluble substrates, we suggest that the insoluble matrix of PLGA could be an obstacle for the interaction with the active centre of the esterases. MmE showed a predominant specificity for fatty esters with short-chain residues [23] which could explain the successful use of the enzyme in the degradation of PLGA. PLGA was not hydrolysed by any of the proteases, thus indicating that co-polyesters of the ResomerÒ series containing glycolic acid residues were apparently not suitable substrates for proteolytic enzymes with esterolytic side activities. It is known from the literature that the presence

Fig. 3. Biodegradability of four ResomerÒ types in the presence of different enzymes (lipases: CaL, CcL, CrL, MmL, PPL, RaL; esterase: MmE; 100 mg/ml each) after 48 h incubation at 37  C. Controls were performed in the same system without enzyme.

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of the protease trypsin could significantly affect the weight loss rate of porous PLGA foams within six weeks [8]. Due to the excessive incubation time, the authors suggested that trypsin may act as a surfactant of the polymer to enhance the dispersion of hydrolytic degradation products into solution but not as a true enzyme. 4. Conclusions The colorimetric-based method described in this paper was successfully validated as a suitable approach for quantitative determination of glycolic acid released as a hydrolysis product of co-polyester PLGA. Advantages of the assay include accuracy, simplicity and low cost. The proposed method clearly demonstrated its efficiency for studying enzymatic hydrolysis of PLGA. Suspensions of PLGA powder with different composition, ResomerÒ RG 503, RG 503 H, RG 653 H and RG 752 H, were biodegraded under controlled conditions applying 22 commercially available hydrolytic enzymes, mainly from microbial origin. Under the conditions employed, seven enzymes (CaL, CcL, CrL, MmL, MmE, PPL and RaL) had significant effects on the polymers with the detection of glycolic acid as indicator of PLGA degradation. Although it is difficult to interpret the observed structural dependence of the enzymatic degradability in terms of a single factor, the co-monomer ratio and the end group of PLGA chains are important factors for the easy approach of the enzyme to the substrate and for the effective enzymatic action. The results reported herein suggest that PLGA co-polyesters are susceptible to enzymecatalysed hydrolysis and that their biodegradability can be widely modified by proper molecular design. Acknowledgements The authors greatly acknowledge funding by the German Ministry of Economics and Technology (BMWi) via grant 16344N from the Federation of Industrial Research Associations (AIF). We are also grateful to Roland Klein (German Institute for Polymers, Darmstadt) for providing Resomer RG 503 and for valuable discussions. Special thanks are extended to Joachim Weber for his excellent assistance in the laboratory. References [1] Y. Tokiwa, B.P. Calabia, Biodegradability and biodegradation of polyesters, J. Polym. Environ. 15 (2007) 259–267.

[2] M. Vert, Aliphatic polyesters: great degradable polymers that cannot do everything, Biomacromolecules 6 (2005) 538–546. [3] A. Södergard, M. Stolt, Properties of lactic acid based polymers and their correlation with composition, Prog. Polym. Sci. 27 (2002) 1123–1163. [4] F. Alexis, Factors affecting the degradation and drug-release mechanism of poly(lactic acid) and poly[(lactic acid)-co-(glycolic acid)], Polym. Int. 54 (2005) 36–46. [5] M. Stevanovic, D. Uskokovic, Poly(lactide-co-glycolide)-based micro and nanoparticles for the controlled drug delivery of vitamins, Curr. Nanosci. 5 (2009) 1–14. [6] Q. Cai, J. Bei, A. Luo, S. Wang, Biodegradation behaviour of poly(lactide-co-glycolide) induced by microorganisms, Polym. Degrad. Stab 71 (2001) 243–251. [7] N. Lucas, C. Bienaime, C. Belloy, M. Queneudex, F. Silvestre, J.-E. Nava-Saucedo, Polymer biodegradation: mechanisms and estimation techniques, Chemosphere 73 (2008) 429–442. [8] Q. Cai, G. Shi, J. Bei, S. Wang, Enzymatic degradation behavior and mechanism of poly(lactide-co-glycolide) foams by trypsin, Biomaterials 24 (2003) 629–638. [9] Y. Tokiwa, B.P. Calabia, Biodegradability and biodegradation of poly(lactide), Appl. Microbiol. Biotechnol. 72 (2006) 244–251. [10] Y. Tokiwa, T. Suzuki, K. Takeda, Hydrolysis of polyesters by Rhizopus arrhizus lipase, Agric. Biol. Chem. 50 (1986) 1323–1325. [11] L.Y. Qiu, In vitro and in vivo degradation study on novel blends composed of polyphosphazene and polyester or polyanhydride, Polym. Int. 51 (2002) 481–487. [12] P. Giunchedi, B. Conti, S. Scalia, U. Conte, In vitro degradation study of polyester microspheres by a new HPLC method for monomer release determination, J. Control. Release 56 (1998) 53–62. [13] F. von Burkersroda, Untersuchung der homogenen und heterogenen Erosion bioabbaubarer Polymere. PhD Thesis, Regensburg, Germany; 2002. [14] A. Niederwieser, A. Matasovic, E.P. Leumann, Glycolic acid in urine. A colorimetric method with values in normal adult controls and in patients with primary hyperoxaluria, Clin. Chim. Acta 89 (1978) 13–23. [15] U. Winkler, M. Stuckmann, Glycogen, hyaluronate, and some other polysaccharides greatly enhance the formation of exolipase by Serratia marcescens, J. Bacteriol. 138 (1979) 663–670. [16] N. Krebsfänger, F. Zocher, J. Altenbuchner, U.T. Bornscheuer, Characterization and enantioselectivity of a recombinant esterase from Pseudomonas fluorescens, Enzym. Microb. Technol. 22 (1998) 641–646. [17] N.J.D. Nagelkerke, A note on a general definition of the coefficient of determination, Biometrika 78 (1991) 691–692. [18] R.A. Jain, The manufacturing techniques of various drug loaded biodegradable poly(lactide-co-glycolide) (PLGA) devices, Biomaterials 21 (2000) 2475–2490. [19] M. Adamczak, Synthesis, properties and application of lipase from Candida antarctica for high yield monoacylglycerol biosynthesis, Pol. J. Food Nutr. Sci. 12 (2003) 3–8. [20] T.G. Park, Degradation of poly(lactic-co-glycolic acid) microspheres: effect of copolymer composition, Biomaterials 16 (1995) 1123–1130. [21] Y. Tokiwa, A. Jarerat, Biodegradation of poly(L-lactide), Biotechnol. Lett. 26 (2004) 771–777. [22] M. Mochizuki, M. Hirami, Structural effects on the biodegradation of aliphatic polyesters, Polym. Advan. Technol. 8 (1997) 203–209. [23] G.J. Moskowitz, R. Cassaigne, I.R. West, T. Shen, L.I. Feldman, Hydrolysis of animal fat and vegetable oil with Mucor miehei esterase. Properties of the enzyme, J. Agric. Food Chem. 25 (1977) 1146–1150.