Hyperbranched polymer conjugated lipase with enhanced activity and stability

Biochemical Engineering Journal 36 (2007) 93–99

Hyperbranched polymer conjugated lipase with enhanced activity and stability Jun Ge, Ming Yan, Diannan Lu, Minlian Zhang, Zheng Liu ∗ Department of Chemical Engineering, Tsinghua University, Beijing 100084, China Received 6 September 2006; received in revised form 20 December 2006; accepted 4 February 2007

Abstract Hyperbranched aromatic polyamides were synthesized from p-phenylenediamine and trimesic acid, and subjected to the conjugation of lipase using carbodiimide as a coupling reagent. The molecular weights of the conjugates were determined by size exclusion chromatography and gel electrophoresis, which showed that each polymer molecule could be covalently modified with a maximum of five to six lipase molecules. The Km of the conjugate to substrate, p-nitrophenylpalmitate, was equal to native lipase while Vmax was increased by 20%, suggesting an enhanced structural transition of lipase during catalysis conducted at the periphery of the polymer. Moreover, the conjugate exhibited a significantly enhanced stability at high temperature or in the presence of organic solvent, as compared to its native counterpart. This might due to the hydrophilic carboxylic groups of polymer that created a suitable microenvironment for lipase. These results indicate that hyperbranched polymer–enzyme conjugate, with the enhanced activity and stability, is promising for industrial biocatalysis. © 2007 Published by Elsevier B.V. Keywords: Bioconjugate; Enzyme modification; Hyperbranched polymer; Hyperbranched aromatic polyamide; Lipase; Enzyme stabilization

1. Introduction Enzyme modification with polymers has been extensively investigated to tailor the various application needs [1], such as “Smart polymers” for enzyme recycling in industrial biocatalysis [2–4], PEG for enhancing enzyme activity in organic solvent [5], polysaccharides or phosopholipid polymers for improving enzyme thermal stability [6,7], and conjugates self-assembly for oil/water biphasic reactions [8]. Preparation of enzyme–polymer conjugates can be achieved by either “grafting-to” or “graftingfrom” methods. In “grafting-to” approaches, polymers are grafted onto the surface of enzyme by reactions between the end groups or pendant groups of polymers and amino acid residues of enzymes. In “grafting-from” approaches, polymer chains are grafted from enzyme surface to fabricate single enzyme nanoparticles [9,10] or by in situ atom transfer radical polymerization (ATRP) to make uniformed enzyme–polymer conjugates [11–13]. By above two approaches, obtained conjugates are given polymer-grafted or crosslinked structures. The surrounding polymer chain, though which is found to be effec-

∗

Corresponding author. Tel.: +86 10 6277 9876; fax: +86 10 6277 0304. E-mail address: [email protected] (Z. Liu).

1369-703X/$ – see front matter © 2007 Published by Elsevier B.V. doi:10.1016/j.bej.2007.02.018

tive in enhancing enzyme functionality and stability, often leads to reduction of the apparent activity of enzyme [5,6,8,9,13], mainly due to the entanglement of polymer chains that cause steric resistance that hinders either the access of enzyme to its substrate and/or the conformational transition of enzyme requested to accomplish the catalysis. Moreover, due to the low reactivity of end groups and pendant groups in polymer, excess amount of polymer should be added in conventional “graftingto” approaches, making the purification of the conjugates more complicated [14]. Conventional linear polymers only have functional groups at one or two ends of the chains, which, moreover, are randomly coiled and entangled with others, resulting in low reactivity of conjugation. Dendritic polymers including dendrimers and hyperbranched polymers [15,16] have a highly branched structure, providing a spherical molecular shape and a high density of functional groups at the periphery. Of particular interest, the large amount of functional chain ends and the globular shape without chain entanglement make the functional groups easier to interact with other reagents, resulting a higher reactivity over the conventional chain polymers [17]. This thus driven a growing attention towards applying dendritic polymers for enzyme immobilization [18,19] and conjugation [20]. With the recognition of the structural advantages of dendritic polymer described

94

J. Ge et al. / Biochemical Engineering Journal 36 (2007) 93–99

above, which make it promising for preparing enzymatic catalyst for industrial applications particular those request high enzyme loading, high activity and high stability, we attempted the preparation of hyperbranched polymer–enzyme conjugate using lipase (EC 3.1.1.3), an enzyme of wide applications in chemicals production in both aqueous and non-aqueous media, such as the decomposition of lipids and the production of cheeselike products, biodiesel and chiral drug intermediates [21,22]. These processes, however, usually conducted at a high temperature and the presence of organic solvents, demand an improved stability of lipase. The present study started with the synthesis of hyperbranched aromatic polyamide by polycondensation between p-phenylenediamine (A2 monomer) and trimesic acid (B3 monomer). Then conjugation of lipase (from Candida rugosa) to the carboxylic groups at the periphery of the polymer was investigated. The composition and the molecular weight of the conjugate were determined by size exclusion chromatography (SEC), and gel electrophoresis. Hydrolysis of p-nitrophenylpalmitate (p-NPP) and p-nitrophenylbutyrate (pNPB) by conjugated lipase was studied and subjected to the comparison to its native counterpart. Finally, the stability of the conjugate and native lipase was determined at high temperature and in the presence of organic solvent.

(KBr, cm−1 ): broad absorption from 3700 to 2000, 1709, 1655, 1516, 1402, 1311, 1244, 831, 669. 2.3. Synthesis of lipase–HBPA conjugate

2. Materials and methods

Protein content in commercial lipase was determined as 11% by bicinchoninic acid (BCA) colorimetric method using bovine serum albumin (BSA) as a standard. In a typical experiment, 100 mg of crude lipase was dissolved in 6 mL of 50 mM, pH 6.5 sodium phosphate buffer containing 50% (v/v) dimethyl sulfoxide (DMSO), followed by centrifugation at 3000 rpm for 5 min. Ten milligrams of hyperbranched aromatic polyamide was dissolved in 400 ␮L of N-methylpyrrolidinone (NMP). Conjugates of different molar ratio of polymer to enzyme were prepared by adding 25, 33, 50 or 100 ␮L of the above polymer solution, respectively, into 1.1 mL of the above enzyme solutions, followed by centrifugation at 3000 rpm for 5 min. Then, 2 mg of 1-ethyl-3-(dimethylaminopropyl) carbodiimide hydrochloride (EDC) was added into 1 mL of the supernatant. After reaction at room temperature for 3 h, the solution was dialyzed with 50 mM, pH 7.0 sodium phosphate buffer for 24 h to remove residual reagents. The unconjugated lipase was removed by ultrafiltartion (MWCO = 100k) using Microcon Centrifugal Filter Devices (Millipore). The amount of conjugated and unconjugated protein was determined by BCA method, according to which the conjugation yield was determined.

2.1. Materials

2.4. Assay

N-Methylpyrrolidinone (NMP) was dried with CaCl2 and then distilled under reduced pressure before use. Pyridine (Py) was dried with KOH, followed by fraction distillation. p-Phenylenediamine (PD), trimesic acid (TMA), triphenyl phosphate (TPP), and 1-ethyl-3-(dimethylaminopropyl) carbodiimide hydrochloride (EDC) of analytical grade were used without further purification. Lipase from C. rugosa (Type VII), p-nitrophenylpalmitate (p-NPP), and p-nitrophenylbutyrate (pNPB) were purchased from Sigma (St. Louis, MO, USA). All the other chemicals were of analytical grade and used without additional treatment.

2.4.1. Reductive SDS-PAGE and agarose gel electrophoresis Reductive SDS-PAGE was performed using 7.5% poly(acrylamide) gels, in which the sample was pretreated with dithiothreitol (DTT). Agarose gel electrophoresis was performed using 7 cm, 1% agarose gels. Forty millimolar, pH 8.3 Tris containing 20 mM sodium acetate and 2 mM EDTA, was used as running buffer and electrophoresis was carried out at the voltage of 100 V for 30 min. Ethidium bromide was used to stain gels after electrophoresis, and then the gels was visualized under UV light. In agarose gel electrophoresis, DNA maker DL 15,000 was used as a standard.

2.2. Synthesis of hyperbranched aromatic polyamide (HBPA) In a typical experiment, 0.25 g (0.0023 mol) of PD, 0.52 g (0.0025 mol) of TMA, 2 mL of Py and 20 mL of NMP were added to a flask with a magnetic stirrer, a nitrogen inlet, and a reflux condenser. The mixture containing 4 mL of TPP and 5 mL of NMP was added dropwise to the solution. After 3 h stirring at 80 ◦ C under N2 atmosphere, the solution was poured into 100 mL of methanol to precipitate the product. The crude product was purified by repeated precipitation from NMP solution into methanol for three times. The product was washed with methanol three times and dried in vacuo at 100 ◦ C. 1 H NMR (500 MHz, DMSO-d6 , δ ppm): 13.46 (br, COOH), 10.66, 10.43 (s, amino proton), 8.80, 8.76, 8.74, 8.71, 8.66, 8.64 (m, aromatic protons from TMA), 7.85, 7.84, 7.82 (m, protons from PD). IR

2.4.2. Size exclusion chromatography Determination of the molecular weight of the conjugate was conducted using size exclusion chromatography using TSKGEL SW4000xL column (TOSHO). The elution buffer was 0.1 M, pH 6.7 sodium phosphate containing 0.1 M Na2 SO4 and 0.05% NaN3 . The experiments were performed at 25 ◦ C and a flow rate of 0.5 mL/min. The calibration curve was established using a kit composed of thyroglobulin (MW: 670,000 g/mol), ferritin (MW: 450,000 g/mol), ␤-galactosidase (MW: 105,000 g/mol), chicken myoglobin (MW: 44,000 g/mol). 2.4.3. Hydrolytic activity assay Specific activity of native and the conjugated lipase was determined by hydrolysis of p-nitrophenylpalmitate (p-NPP) and p-nitrophenylbutyrate (p-NPB) using a standard method

J. Ge et al. / Biochemical Engineering Journal 36 (2007) 93–99

[23]. The p-nitrophenylesters were pre-dissolved in acetonitrile and then diluted into sodium phosphate buffer (50 mM, pH 7.0) with desired concentration and followed by sonication for 3 min. The reaction was started by adding 50 ␮L of enzyme solution (50 ␮g/mL) to the 950 ␮L substrate solution and followed by measurement of absorbance changes at 348 nm for 2–5 min. The acetonitrile concentration in the reaction solution was maintained at 5% (v/v). The molar extinction coefficient in this condition calculated by calibration curve is 4700 M−1 cm−1 . And, during the assay, the conjugate solution concentration was determined by BCA method based on enzyme. Triton X-100 was added in the p-NPP solution in the lipase activity assay to enhance the solubility of p-NPP. In order to minimize the either the enhancement or inhibition of Triton X-100 on the lipase activity, as reported elsewhere [24], the concentration of Triton X-100 was maintained at 0.02% (v/v), well above the CMC of Trition X-100 (0.015%). Release of 1 ␮mol of p-nitrophenol per minute in the assay conditions is defined as one unit activity (U mg−1 ). 2.4.4. Measurements 1 H NMR spectra were recorded on a JEOL JNM-ECA600 NMR spectrometer. Infrared spectra were recorded using a Nicolet 560-IR Fourier transform infrared spectrometer. UV–vis spectrum were recorded on SHIMADZU MultiSpec-1501 UV–vis spectrometer. 3. Results and discussion 3.1. Characterization of hyperbranched aromatic polyamide (HBPA) The synthesis of hyperbranched aromatic polyamide developed by Kakimoto [25] is shown in Scheme 1. The hyperbranched aromatic polyamide was obtained by polycondensation of p-phenylenediamine (PD) and trimesic acid (TMA) using triphenyl phosphate (TPP) and pyridine (Py) as condensing agents. The integration ratio for the peaks assigned to aromatic protons of the PD and TMA units was determined as

95

1.4:1.0. Thus molar ratio of PD and TMA units incorporated in polymer was interpreted as 0.35:0.33. Multiple peaks from 8.80 to 8.64 ppm were assigned to the protons of all 1,3,5trisubstituted benzene moieties including linear (L), dendritic (D), and terminal (T) units (as shown in an enlarged region in Fig. 1). The signals at 8.80, 8.66 ppm were assigned to the protons from structure 1 and structure 2, respectively. Structrue 1 and 2 are linear units [26]. By integration ratio for the peaks attributed to aromatic protons of linear unit and of all 1,3,5trisubstituted benzene moieties, the degree of branching (DB) calculated according to the equation: DB = (D + T)/(D + L + T) [27] was 0.54. As shown in Fig. 2, the absorption at 1709 cm−1 suggested the existence of carboxylic groups, and the absorption at 1655, 1554 cm−1 suggested the formation of the amide. The state of the hyperbranched aromatic polyamide in aqueous media was determined by agarose gel electrophoresis described above. The apparent molecular weight of hyperbranched polymer evaluated by agarose gel electrophoresis using DNA as a standard was about 150,000 g/mol (1 bp ≈ 500 Da), as shown in Fig. 5b. 3.2. Synthesis and characterization of lipase–HBPA conjugate The size exclusion chromatography of the purified conjugate and native lipase is shown in Fig. 3. Here the conjugate peaks is excluded earlier than the native lipase, indicating a major difference in molecular weight and size. The mixture of lipase and hyperbranched aromatic polyamide in absence of coupling reagent did not show a similar peak that can be designed to the conjugate. As shown in Fig. 4a, with the increase in the feed ratio of polymer to lipase in synthesis of conjugates 1, 2, 3 and 4 (molar ratio of lipase to polymer = 8:1, 6:1, 4:1 and 2:1), the retention volume of conjugates 1–4 in SEC curves increases gradually, indicating a gradually decrease in molecular weight of conjugates. As shown in Fig. 4b, calculated from calibration curve, the molecular weight of conjugates 1, 2, 3 and 4 is 468,000, 459,000, 363,000 and 250,000 g/mol, and the

Scheme 1. Preparation of hyperbranched polymer conjugated lipase.

96

J. Ge et al. / Biochemical Engineering Journal 36 (2007) 93–99

Fig. 1. 1 H NMR spectrum of hyperbranched aromatic polyamide in DMSO-d6 .

number of lipase molecules bounded to each polymer is 5.6, 5.4, 3.7 and 1.8, respectively. This suggested that the conjugate possesses a structure in which each polymer molecule is covalently modified with several enzyme molecules, rather than a polymer-grafted or crosslinked structure in which molecular weight of conjugates should increase more dramatically with the increase in feed ratio of polymer to enzyme. Note that the number of lipase in conjugate 1 and 2, being 5.6 and 5.4, respectively, approaches to each other, indicating a possible maximum number of lipase molecules accommodated by the polymer. Moreover, the conversion of lipase in conjugate 1 was 65% at a molar ratio of lipase to polymer at 8:1, while the conversation at a lower molar ratio reaches 80–90%, (conjugates 2, 3 and 4). This also indicates the possible maximum accommodation of lipase molecules by the dendritic polymer. The hyperbranched poly-

Fig. 2. IR spectrum of hyperbranched aromatic polyamide.

mer has a diameter ranging between 10 and 50 nm according to TEM measurement while lipase molecule has a dimension of 6.88 nm × 6.85 nm × 5.2 nm (data from PDB, website: http://www.rcsb.org/pdb/explore.do?structureId=1TRH). In addition to the size effect, the maximum number of enzyme

Fig. 3. Size exclusion chromatography of separated conjugate, native lipase. The experiments were performed as described in Section 2.

J. Ge et al. / Biochemical Engineering Journal 36 (2007) 93–99

97

3.3. Hydrolytic activity of native lipase and conjugate

Fig. 4. (a) Size exclusion chromatography of bioconjugate product with different molecular weight; (b) the molecular weight of conjugate vs. the concentration of polymer added into the same reaction system. The experiments were performed as described in Section 2 (1: conjugate 1, 2: conjugate 2, 3: conjugate 3, 4: conjugate 4).

conjugated onto polymer is determined by the density of carboxyl groups at the periphery of polymer. Fig. 5a shows the reductive SDS-PAGE of the native lipase and the conjugate 1, in which the conjugate accumulated at the top of the gel. Whereas Fig. 5b is the agarose gel electrophoresis of conjugate 1, from which the molecular weight of conjugate 1 was interpreted as 500,000 g/mol (1 bp ≈ 500 Da). The number of lipase in each conjugate was 6.1, similar to that determined by SEC.

Michaelis–Menten equation was applied to the hydrolysis of p-nitrophenylpalmitate (p-NPP) and p-nitrophenylbutyrate (pNPB) by HBPA conjugated (conjugate 1) and native lipase, respectively. In the experiment, the concentration of p-NPP and p-NPB was increased from 0 to 1.0 mM. Km and Vmax interpreted from Lineweaver–Burk plot are listed in Table 1. The increase in Km , due to extra mass transport resistance by the matrix on which enzyme was immobilized is often observed in the conventional enzyme immobilization [28]. For the present study, however, Km for native lipase and conjugate is almost identical, i.e., the conjugation does not lead to steric hindrances for the access of substrate. This can be predicated from the proposed structure of the conjugate (shown in Scheme 1) that, the existence of lipase at the surface of the hyperbranched polymer that enables a similar access to substrate, as compared to the native lipase in solution. It is interesting to note that Vmax of conjugate is 120% as much as that of native lipase for hydrolysis of p-NPP, and 110% for hydrolysis of p-NPB, indicating the enhancement of the enzymatic reaction. Interfacial activation is one established mechanism for lipase, and an “open” conformation with high activity is generated once lipase is adsorbed to the hydrophobic interface [21]. Some researches [29,30] reported that lipase adsorbed on some supports exhibits enhanced hydrolytic activity. For the present study, lipase was bound to the periphery of hyperbranched polymer which has a hydrophobic core. These special surface properties may also favorable for the interfacial activation of lipase consequently. 3.4. Thermal stability of native lipase and conjugate The stability of conjugated lipase (conjugate 1) and native lipase at 50 and 80 ◦ C was determined in 50 mM, pH 7.0 sodium phosphate buffer with the same protein concentration

Fig. 5. (a) SDS-poly(acrylamide) gel electrophoresis of native lipase and lipase conjugate (lane 1: native lipase, lane 2: lipase conjugate, lane 3: protein marker); (b) Agarose gel electrophoresis of lipase conjugate and hyperbranched aromatic polyamide (lane 1: DNA marker, lane 2: lipase conjugate, lane 3: polymer).

98

J. Ge et al. / Biochemical Engineering Journal 36 (2007) 93–99

Table 1 Activity and kinetic parameters of native lipase and lipase conjugate p-NPP

Km (mM) Vmax (U mg−1 )

p-NPB

Native lipase

Lipase conjugate

Native lipase

Lipase conjugate

0.25 26 (100%)a

0.24 31 (120%)

0.28 87 (100%)a

0.25 95 (110%)

a Commercial lipase from Sigma contains two isoenzymes: lip1 (89%), lip3 (11%). Specific activity of two isoenzymes: lip1 has activity of 12.0 U mg−1 for hydrolysis of p-NPP, and 50.9 U mg−1 for hydrolysis of p-NPB; lip3 has activity of 222 U mg−1 for hydrolysis of p-NPP, and 351 U mg−1 for hydrolysis of p-NPB [23].

of 50.0 ␮g/mL. Aliquots were withdrawn periodically from test tubes in water bath and then subjected to the measurement of hydrolytic activity as described in Section 2. Assuming the thermal inactivation follows a two-step series model [31], the experimental plots of relative activity versus incubation time were adjusted to exponential decay, as shown in Fig. 6. At 50 ◦ C, the conjugate has higher stability than native one, and the half-life of the enzyme activity was extended from 55 min (native form) to 180 min (conjugated form). And, at 80 ◦ C, native lipase was deactivated within 30 min, while it took 90 min to fully deactivate the conjugates. The enhanced thermal stability after immobilization was reported by many other researchers [28,32]. The existence of the multiple covalent bonds between polymer and enzyme was the major reason leading to an enhanced stability by preventing protein unfolding at high temperature, whereas native one in solution loses its structure due to intensified thermo fluctuation at high temperature. 3.5. Stability of native lipase and conjugate in organic solvent The exposure to organic solvent leads to the denaturation of many enzymes mainly due to the loose of ‘essential water’ bounded at some critical regions of enzyme which is necessary for enzyme to the display its biological function [33,34]. For present study, DMSO was applied to examine the stability of lipase in non-aqueous enzymatic catalysis [22]. In order to observe the irreversible deactivation, native lipase and the

Fig. 7. Comparing of residual activities of native lipase and the conjugate with the same protein concentration (25 ␮g/mL) incubated in 50 mM, pH 7.0 sodium phosphate buffer containing DMSO with different concentration at 40 ◦ C for 20 h.

conjugate (conjugate 1) were incubated in organic media for a period of time, followed by the measurement of residual hydrolytic activity in aqueous solution [35] using a standard method as described in Section 2. The residual activities of native lipase and the conjugate after 20 h incubation in 50 mM, pH 7.0 sodium phosphate buffer containing 10, 20, 30, 40% (v/v) of DMSO at 40 ◦ C are given in Fig. 7. The native lipase and the conjugate were incubated with the same protein concentration of 25.0 ␮g/mL. Here, native lipase remained 49, 44, 38, 27% of its initial activity while conjugated lipase remained 99, 89, 73, 50%, respectively. It has been reported that enzyme located at the hydrophilic environment provided by carboxylic groups exhibits an increased stability in organic solvent [36]. For hyperbranched aromatic polyamide conjugated lipase, the significantly enhanced stability may be attributed to the large amount of carboxylic groups at the polymer’s periphery, which provide a hydrophilic microenvironment that enables lipase to maintain the “essential water” for its biological function. 4. Conclusions

Fig. 6. Thermal inactivation of native lipase and the conjugate with the same protein concentration (50 ␮g/mL) in 50 mM, pH 7.0 sodium phosphate buffer at 50 and 80 ◦ C.

A hyperbranched aromatic polyamide (HBPA) was synthesized using polycondensation between p-phenylenediamine and trimesic acid. Conjugation of lipase to the carboxylic groups of the HBPA was accomplished in presence of coupling reagent. The size exclusion chromatography and gel electrophorsis suggested that each HBPA molecule could be covalently modified

J. Ge et al. / Biochemical Engineering Journal 36 (2007) 93–99

with a maximum of five to six lipase molecules. The conjugated lipase had an approximate Km to the native lipase while Vmax is increased by 20% for the hydrolysis of p-nitrophenylpalmitate (p-NPP). Moreover, the conjugated lipase exhibited a significantly higher stability at high temperature or in the presence of DMSO. It is excepted from above experimental results that conjugation of lipase to the HBPA enables its free contact to substrate while the sounding carboxyl groups located the surface of HBPA creates a hydrophilic microenvironment that enables the conjugated lipase to display biological activity at high temperature or in the presence of organic solvent. These properties of HBPA–lipase conjugate make it suitable for potential industrial application. Acknowledgment This work was supported by the National Natural Science Foundation under Grant No. 2033601. References [1] G. DeSantis, J.B. Jones, Curr. Opin. Biotechnol. 10 (1999) 324. [2] Z.L. Ding, G.H. Chen, A.S. Hoffman, Bioconjugate Chem. 7 (1996) 121. [3] Y. Ito, N. Sugimura, O.H. Kwon, Y. Imanishi, Nat. Biotechnol. 17 (1999) 73. [4] M. Yan, J. Ge, W. Dong, Z. Liu, P. Ouyang, Biochem. Eng. J. 30 (2006) 48. [5] A. Matsushima, Y. Kodera, M. Hiroto, H. Nishimura, Y.J. Inada, J. Mol. Catal. B: Enzyme 2 (1996) 1. [6] R.M. de la Casa, J.M. Guis´an, J.M. S´anchez-Montero, J.V. Sinisterra, Enzyme Microb. Technol. 30 (2002) 30. [7] D. Miyamoto, J. Watanabe, K.J. Ishihara, Appl. Polym. Sci. 95 (2005) 615. [8] G. Zhu, P. Wang, J. Am. Chem. Soc. 126 (2004) 11132. [9] J. Kim, J.W. Grate, Nano Lett. 3 (2003) 1219.

99

[10] M. Yan, J. Ge, Z. Liu, P. Ouyang, J. Am. Chem. Soc. 128 (2006) 11008. [11] K.L. Heredia, D. Bontempo, T. Ly, J.T. Byers, S. Halstenberg, H.D. Maynard, J. Am. Chem. Soc. 127 (2005) 16955. [12] D. Bontempo, H.D. Maynard, J. Am. Chem. Soc. 127 (2005) 6508. [13] B.S. Lele, H. Murata, K. Matyjaszewski, A.J. Russell, Biomacromolecules 6 (2005) 3380. [14] C.J. Fee, J.M. Van Alstine, Chem. Eng. Sci. 61 (2006) 924. [15] Y.H. Kim, O.W. Webster, J. Am. Chem. Soc. 112 (1990) 4592. [16] C. Gao, D. Yan, Prog. Polym. Sci. 29 (2004) 183. [17] J.M. Fr´echet, Science 263 (1994) 1710. [18] O. Yemul, T. Imae, Biomacromolecules 6 (2005) 2809. [19] M.E. Cosulich, S. Russo, S. Pasquale, A. Mariani, Polymer 41 (2000) 4951. [20] G. Chen, D. Huynh, P.L. Felgner, Z. Guan, J. Am. Chem. Soc. 128 (2006) 4298. [21] R.D. Schmid, R. Verger, Angew. Chem. Int. Ed. 37 (1998) 1608. [22] G. Carrea, S. Riva, Angew. Chem. Int. Ed. 39 (2000) 2226. [23] N. L´opez, M.A. Pernas, L.M. Pastrana, A. S´anchez, F. Valero, M.L. R´ua, Biotechnol. Prog. 20 (2004) 65. [24] P. Helist¨o, T. Korpela, Enzyme Microb. Technol. 22 (1998) 113. [25] M. Jikei, S. Chon, M. Kakimoto, S. Kawauchi, T. Imase, J. Watanebe, Macromolecules 32 (1999) 2061. [26] H. Komber, B. Voit, O. Monticelli, S. Russo, Macromolecules 34 (2001) 5487. [27] C.J. Hawker, R. Lee, J.M.J. Fr´echet, J. Am. Chem. Soc. 113 (1991) 4583. [28] S.H. Chiou, W.T. Wu, Biomaterials 25 (2004) 197. [29] A. Bastida, P. Sabuquillo, P. Armisen, R. Fern´andez-Lafuente, J. Huguet, J.M. Guis´an, Biotechnol. Bioeng. 58 (1998) 486. [30] L. Yu, I.A. Banerjee, X. Gao, N. Nuraje, H. Matsui, Bioconjugate Chem. 16 (2005) 1484. [31] J.P. Henley, A. Sadana, Enzyme Microb. Technol. 7 (1985) 50. [32] J.M. Moreno, M. Arroyo, M.J. Hern´aiz, J.V. Sinisterra, Enzyme Microb. Technol. 21 (1997) 552. [33] B. Schulze, A.M. Klibanov, Biotechnol. Bioeng. 38 (1991) 1001. [34] L.A.S. Gorman, J.S. Dordick, Biotechnol. Bioeng. 39 (1992) 392. [35] M.N. Gupta, R. Batra, R. Tyagi, A. Sharma, Biotechnol. Prog. 13 (1997) 284. [36] H. Chen, Y.L. Hsieh, Biotechnol. Bioeng. 90 (2005) 405.