Water-Stable All-Biodegradable Microparticles in Nanofibers by

Article pubs.acs.org/Biomac

Water-Stable All-Biodegradable Microparticles in Nanofibers by Electrospinning of Aqueous Dispersions for Biotechnical Plant Protection Priyanka Bansal, Kathrin Bubel, Seema Agarwal, and Andreas Greiner* Philipps-Universität Marburg, Department of Chemistry and Scientific Centre for Materials Science, Hans-Meerwein-Strasse, Geb. H, D-35032 Marburg, Germany ABSTRACT: Pheromone eluting oligolactide (OLA) microcapsules immobilized in electrospun biodegradable polyester nanofibers were obtained by electrospinning of aqueous dispersions of the microcapsules. OLA was prepared by conventional melt polycondensation of lactic acid. Following the protocol of the solvent displacement method, OLA was dissolved in acetone and mixed with Brij S20 and the pheromone of the European grape vine moth, Lobesia Botrana, (E,Z)-7,9-dodecadien-l-yl acetate (DA). Up to 32 wt % of this mixture could be dispersed in water with colloidal stability of several weeks without any sedimentation. Without DA as well as OLA, no stable dispersions of OLA in water were obtained. Replacement of DA by classical hydrophobes typically used for miniemulsions did not yield stable dispersions, but the addition of octyl acetate, which shows structural similarity to DA, yielded stable dispersions in water up to 10 wt %. Dispersions of OLA/DA were successfully electrospun in combination with an aqueous dispersion of a biodegradable block copolyester resulting in water-stable nanofibers containing OLA/DA microcapsules. Release of DA from microcapsules and fibers was retarded in comparison with non-encapsulated DA, as shown by model studies.

■

INTRODUCTION Biotechnical plant protection has been achieved successfully by release of female pheromones from artificial dispensers, leading to confusion of male insects and in consequence to mating disruption.1,2 However, this approach poses a problem in maintaining a sufficiently high and homogeneously distributed concentration of the pheromone in the field for a longer period. Two methods are being extensively employed to overcome this problem. One is to place large dispensers filled with the pheromone at fixed points in the treatment areas.3−6 The other way is by microencapsulating the pheromone, which is then directly applied to the crop by conventional methods.7−10 Hence an approach is needed that (i) makes the pheromone less prone to being swept away by storms and heavy rains, (ii) controls the rate at which they are released, and (iii) causes a homogeneous distribution of the pheromone over the field, cutting down the large amount of the pheromone otherwise needed in case of dispensers. A promising concept by use of pheromone-loaded electrospun polymer nanofiber-based nonwovens has been recently suggested.11 Major drawbacks for electrospinning in agricultural environment are the need for harmful solvents for electrospinning of biodegradable water stable nonwovens and the uncontrolled and fast release of pheromones. Recently, electrospinning of aqueous polymer suspensions has been established as highly versatile approach for the formation of water stable nonwovens.12−16 Biodegradable but water-stable nanofibers nonwovens were obtained by electrospinning of concentrated aqueous dispersions of block copolyesters.15 Waterstable composite nanofibers with biodegradable polyester © 2012 American Chemical Society

nanoparticles and an encapsulated dye as a model drug were obtained by electrospinning of PVA and suspended poly(lactideco-glycolide) nanoparticles.16 On the basis of this, we wondered whether biodegradable pheromone-loaded microparticles taking a large amount of pheromone could be dispersed in water and could be electrospun along with the block copolyesters previously used for electrospinning.15 Our concept was to disperse OLAs in water as shell material for pheromone microcapsules. OLAs are well-known to be biodegradable but not dispersible in large quantities in water. Our initial assumption was that highly concentrated dispersions of OLAs could be stabilized in water as a dispersion by the ionic end groups of the OLA chain giving a tenside effect. This assumption turned out to be wrong in the course of our investigations. Interestingly, we found that the pheromone itself as well as structurally similar compounds stabilize the aqueous dispersion of OLA microparticles. The results of these findings as well as our attempts to prepare microparticle-loaded block copolyester nonwovens from aqueous formulations by electrospinning is reported here.

■

EXPERIMENTAL SECTION

Materials. D,L-Lactic acid (Aldrich) was purified by distillation. (E,Z)-7,9-Dodecadien-l-yl acetate (DA) (Trifolio-M), Brij S20/Brij 78 (Aldrich), PEO (Mw = 900 000 Da; Acros Organics), and PVA (Mowiol 8-88) (Aldrich) were used as received. Acetone was distilled prior to use. Received: October 19, 2011 Revised: December 27, 2011 Published: January 3, 2012 439

dx.doi.org/10.1021/bm2014679 | Biomacromolecules 2012, 13, 439−444

Biomacromolecules

Article

Measurements and Equipment. 1H and 13C NMR spectroscopy was done with a Bruker ARX400 spectrometer in CDCl3 solution and the software MestRec version 4.9.9.6. Thermogravimetrical decomposition was analyzed by an 851 TG module from Mettler in a nitrogen atmosphere (flow rate: 50 mL/min). We placed 10−12 mg of the sample in an alumina crucible, which was heated isothermally to 30 °C for 4 h. Differential scanning calorimetry (DSC) was done by a Mettler Toledo DSC 821c using heating/cooling rates of 10 K/min under a N2 atmosphere. Evaluation was done with the second heating run and STARe software. Particle size analysis was carried out by dynamic light scattering using a Beckman Coulter Delsa Nano C particle size analyzer. The average particle diameters were found from the cumulant results. Ultrasound treatment was performed with an ultrasonic device (Bandelin electronic UW 60, Bandelin Sonoplus HD 60 adapter, power 70 W). For dialysis, dialysis membrane Spectra Por (MWCO = 1000; normal width = 38 mm, diameter = 24 mm) was used. The dialysis tubes were precleaned by treatment with deionized water for 20 min. Electrospinning was done using a previously described setup.17 Synthesis of OLA. We weighed 200 g of D,L-lactic acid (90% in water) in a precleaned and dried 500 mL round-bottomed flask. The temperature was increased to 100 °C to distill off the 10% water present in the monomer without applying vacuum to prevent distillation of D,L-lactic acid (boiling point = 122 °C @ 12 mmHg). The heating was then increased stepwise to 200 °C simultaneously removing water by vacuum distillation. Dispersions of OLA/Brij S20/DA in Water. For the preparation of 1 wt % dispersion in water, 0.1 g of OLA was dissolved in 2.5 mL of acetone. After the addition of a solution containing 10 mL of deionized water, 0.1 g Brij S20, and 0.1 g of DA, the mixture was subjected to ultrasound for 4 min. The resulting dispersion was purged under a mild air stream at 20 °C to remove acetone. For the preparation of 10 wt % dispersion in water, 1 g of OLA was dissolved in 5 mL of acetone. After the addition of a solution containing 10 mL of water, 0.1 g Brij S20, and 1 g of DA, the mixture was subjected to ultrasound for 4 min. The resulting dispersion was purged under mild air stream at 20 °C to remove acetone. Preparation of OLA Dispersions by Up-Concentration via Dialysis of OLA Dispersions. A larger batch of a new aqueous dispersion of OLA (10 wt % of OLA/Brij S20/DA, Mn ≈ 954 Da, particle size = 478 nm) was used for the up-concentration by dialysis. We filled 50 mL of the 10 wt % dispersion in the dialysis tube with a length of 10 cm and placed it in 800 mL of an aqueous solution of 15 wt % PVA (Mw = 14 000 Da) for 72 h at room temperature. After 72 h, the tube was removed from the PVA solution and rinsed with water and dried. Solid content of OLA was obtained by freeze-drying and weighing. Synthesis of PHA-b-MPEG Dispersion (2.5 wt %) in Water. The synthesis of the diblock copolyester was performed by melt polycondensation using adipic acid, 1,6-hexanediol, and α-hydroxy-ωmethoxy-PEO (Mw = 5000 Da) previously reported.15 In short: 0.5 g of PHA-b-MPEG (Mn = 6400 Da, PDI = 2.1) was dissolved in 12.5 mL of acetone. After the addition of a solution of 20 mL of water and 0.05 g of Brij 78, the mixture was subjected to ultrasound for 4 min. The resulting suspension was purged under a mild air stream at 20 °C to remove acetone. Upconcentration of 2.5 wt % dispersion of PHA-b-MPEG (Mn = 6400 Da, PDI = 2.1, particle size = 108 nm) to 16 wt % was carried out by dialysis. 500 mL of the dispersion was filled in a dialysis tube with a length of 15 cm and placed in 6 L of an aqueous solution of PVA (15 wt %). After 100 h, the tube was removed from the PVA solution and rinsed with water. The solid content of PHA-b-MPEG was obtained by freeze-drying and weighing. Preparation of Samples for Release Study of the Pheromone. Two solutions were prepared to study the release rate of the pheromone. The first solution was prepared by mixing 5 g of 32 wt % OLA/pheromone/Brij S20 dispersion (prepared by a combination of solvent displacement method and dialysis), 5 g of 2.5 wt % PHA-b-MPEG dispersion, and 0.4 g of PEO (Mw = 900 000 Da) as the template polymer.

The other solution was prepared by the addition of 5 g of 2.5 wt % PHA-b-MPEG dispersion, 0.4 g of PEO (Mw = 900 000 Da), and 1 g of free pheromone. Sample films for analysis were prepared from both the solutions by solvent casting method. In addition, pheromone release was studied on electrospun nonwovens by isothermal thermogravimetrical analysis at 30 °C for 4 h. Electrospinning of OLA/DA in PHA-b-MPEG with PEO. We mixed 1 g of 1 wt % OLA/pheromone/Brij S20 dispersion with 4 g of 16 wt % PHA-b-MPEG dispersion. We added 0.2 g of PEO (Mw = 900 000 Da) to this mixture. Electrospinning was carried out by applying a feed of 0.05 mL/min and an electrode distance of 22 cm at a voltage of 15 kV. Aluminum foil was used as a substrate for collecting the electrospun fibers.

■

RESULTS AND DISCUSSION OLA Synthesis and Characterization. OLA was prepared by polycondensation of D,L-lactic acid at a temperature starting from 100 to 200 °C stepwise for a time of 16 h by simultaneous elimination of water under vacuum (Scheme 1). Scheme 1. Synthesis of OLA by Polycondensation of D,LLactic Acid

OLA was characterized by 1H NMR spectroscopy. A ratio of 12:1 was calculated by comparison of the integrals of the peaks at δ = 5 ppm (−O−CH(CH3)−C(O)− of repeat unit) and δ = 4.2 ppm (HO−CH(CH3)−C(O)− of terminal unit), which implied that there were 13 repeating units in the OLA, which amounted to Mn ≈ 950 Da. Thermal analysis of bulk OLA by DSC showed a Tg of 12−15 °C, which confirmed the macroscopic waxy appearances. Preparation and Characterization of OLA Microparticle Dispersions. The OLA was dissolved in acetone. After the addition of a solution of water and Brij S20, ultrasound was applied. Finally, acetone was removed by slow evaporation under mild air stream at 20 °C. Following this procedure, precipitation of OLA occurred with OLA as an oily residue on the water surface (Figure 1A). The dispersion of

Figure 1. Photos of (A) unstable OLA without DA and (B) a stable aqueous dispersion by solvent displacement method of 10% OLA, 10% DA, and 1% Brij S20 after ultrasonification and removal of acetone and (C) unstable 10 wt % aqueous dispersion containing DA/Brij S20 without OLA.

OLA and Brij S20 together with DA resulted in stable milky dispersions without any visible agglomeration or segregation of 440

dx.doi.org/10.1021/bm2014679 | Biomacromolecules 2012, 13, 439−444

Biomacromolecules

Article

OLA on the water surface (Figure 1B) with an average particle size of 0.112 to 1.2 μm (Figure 2). In contrast, 10 wt % aqueous

Table 2. Stability and Average Particle Size of 1 wt % Aqueous Dispersions of OLA and Brij S20 as a Function of the Different Additives Replacing DA (OLA:DA is 1:1)

Figure 2. Particle size versus intensity (%) by dynamic light scattering of 10 wt % aqueous dispersion of OLA/DA/Brij S20.

showed structural analogy to the DA, yielded dispersions with similar results like DA. Stable dispersions up to 10 wt % were observed by using OA in place of DA, as shown in Figure 3.

dispersions of DA/Brij S20 without the addition of OLA were also unstable (Figure 1C). Obviously, the pheromone DA plays an important role in the formation of stable aqueous dispersions of OLA. Therefore, the stability of the dispersions and the average particle size were investigated using different concentrations of DA in 1 wt % aqueous dispersions (Table 1). Table 1. Stability and Average Particle Size of 1 wt % Aqueous Dispersions of OLA and Brij S20 as a Function Content of DA S. No.

OLA:DA

stability of the dispersion

average diameter/nm

1 2 3 4 5 6

1:0.2 1:0.3 1:0.5 1:0.7 1:0.9 1:1

instable instable stable stable stable stable

164 185 285 321 357 408

Table 1 shows that the average particle size of the dispersion increases with increase in the concentration of DA. However, the dispersions containing DA in 20 and 30 wt % ratios were found to be highly unstable. The amount of DA required to stabilize the dispersion was found to be ∼50 wt %, which showed a monomodal particle size distribution centered around 408 nm. It is obvious so far that the hydrophobic DA plays a major role for the stabilization of OLA microparticles in aqueous dispersions. It is well known in miniemulsion that a hydrophobe compound can prevent aggregation of emulsions. Therefore, we wondered whether DA could be replaced by other hydrophobes, for example, hexadecane, which is wellestablished for the stabilization of miniemulsions.18 To investigate the influence of different hydrophobes, we added each of these hydrophobes to prepare 1 wt % dispersion, and the particle size and the stability were compared (Table 2). Hydrophobic compounds like hexadecane and cetyl alcohol (hexadecanol), which are typically used as hydrophobes in miniemulsions, yielded dispersions with lower stability. In addition, most of the hydrophobes yielded dispersions with large particle diameters. However, octyl acetate (OA), which

Figure 3. (A) 1 wt % OLA dispersion and (B) 10 wt % OLA dispersion stabilized by the addition of OA replacing DA.

Analysis of the particle diameter by dynamic light scattering showed an average diameter of 238 nm for 1 wt % OLA/OA/ Brij S20 dispersion and 400 nm for 10 wt % OLA/OA/Brij S20 dispersion and a monomodal particle size distribution. However, the solid content of the dispersions prepared could be increased drastically without aggregation by osmosis similar to the procedure previously described.15 Here a dialysis tube containing a 10 wt % aqueous dispersion of OLA, DA, and Brij S20 was placed in a PVA solution (15 wt % in water). The dialysis was stopped after 72 h to avoid agglomeration. After 72 h of dialysis, stable aqueous dispersions of OLA up to 32 wt % with an average particle size of ∼450 nm remained in the dialysis tube. This 32 wt % aqueous dispersion of OLA/DA tenside showed remarkable colloidal stability in water to at least 2 weeks. Furthermore, for DA to be present in the OLA microparticles, it is important that it is miscible with the OLA. The miscibility of 441

dx.doi.org/10.1021/bm2014679 | Biomacromolecules 2012, 13, 439−444

Biomacromolecules

Article

pheromone in OLA was proven by DSC analysis of a mixture containing 34 wt % DA with respect to OLA. The DSC analysis was performed from a temperature of −50 to 50 °C at a heating rate of 10 °C/min (Figure 4).

Figure 4. DSC of pure OLA and OLA microcapsules loaded with 34 wt % of DA.

From the DSC curves, the glass-transition temperature of the OLA was found to be 12 °C. On mixing 34 wt % pheromone with OLA, the glass-transition temperature of the OLA decreased to 9 °C. This showed that the pheromone acted as plasticizer and indicated the miscibility of the pheromone in OLA and hence the presence of pheromone in the OLA microparticles. The effect of microencapsulation on the volatility of the pheromone was studied by TGA and will be discussed below in combination with release studies of the pheromone from electrospun fibers. Electrospinning of OLA/DA in PHA-b-MPEG with PEO. To immobilize OLA/DA microparticles in biodegradable electrospun nonwovens, electrospinning of 1 wt % OLA/Brij S20/ DA dispersion was carried out with a 16 wt % water-based dispersion of PHA and α-hydroxy-ω-methoxy-PEG (Mn = 5000 Da) diblock copolyester (Figure 5). The PHA-b-MPEG dispersion

Figure 6. Intensity distribution of particle size obtained by DLS analysis of the 1 wt % OLA/Brij S20/DA (top), 16 wt % PHA-bMPEG dispersion (middle), and mixed dispersion of 1 g of the 1 wt % OLA/Brij S20/DA dispersion and 4 g of the PHA-b-MPEG dispersion as used for electrospinning (lower).

this state, we cannot rule out that the particles of both dispersions merged together to form mixed particles. Cylindrical bead-free fibers were obtained by electrospinning of the mixed dispersion with PEO as viscosity modifier (Figure 7A)

Figure 5. Structure of Polyhexyleneadipate-block-methoxypolyethyleneglycol (PHA-b-MPEG).

was used owing to its high solid content and good electrospinnability. High-molecular-weight PEO (Mw = 900 000 Da) was used as a viscosity modifier, as previously published.15 To study the effect on particle size by combination of dispersions of the pheromone containing OLA microparticles and the PHA-b-MPEG dispersion, we performed DLS analysis (Figure 6). The reproduction of the 1 wt % OLA/Brij S20/DA showed nice reproduction of the average particle size here of 408 nm (compare Table 1, S. No. 6 = 400 nm), whereas the particle size of the 16 wt % PHA-MPEG dispersion was found to be centered around 118 nm, in good agreement with the previously reported value.15 The particle size distribution of the mixed dispersion showed an average particle size of 194 nm. In

Figure 7. SEM images of electrospun fibers of 1 wt % OLA/Brij S20/DA aqueous dispersion with PHA-b-MPEG dispersion and PEO (A) before water treatment and (B) after water treatment for 24 h at 20 °C.

with average fiber diameter of 500 nm. Obviously the dispersion particles merged together by fiber formation, as previously observed for electrospinning of acrylate dispersions.13 After electrospinning, the fibers were treated with water to remove PEO and to probe the water stability of the fibers. The fibers remained intact despite water treatment. The fiber diameter did not change significantly after removal of PEO (analyzed by IR 442

dx.doi.org/10.1021/bm2014679 | Biomacromolecules 2012, 13, 439−444

Biomacromolecules

Article

and NMR spectroscopy, not shown here), but the fiber surface became uneven, as it can be seen in the inset of Figure 7B. The release of the pheromone DA from electrospun fibers before and after water treatment was analyzed by isothermal thermogravimetrical analysis at 30 °C and compared with the volatility of free DA and DA in microparticles (Figure 8).

release of pheromone is required for 8 weeks. The fibers produced here by electrospinning contained ∼1.1 wt % of DA, which means according to the TGA measurements all pheromone is released after ∼7 h (for fibers before water treatment), which is clearly too fast for the real world application.

■

CONCLUSIONS Biodegradable OLAs were successfully dispersed in water to concentrations of ∼10 wt %. The addition of DA in combination with Brij S20 to OLA resulted in aqueous dispersion of a total solid content (OLA, DA, Brij S20) of up to 32 wt % applying the combination of solvent displacement method and dialysis. The OLA/DA dispersions were stable for several weeks without any sedimentation. No stable dispersion of OLA in water was possible without DA as well as no stable dispersions of DA were possible without OLA. The minimum DA concentration required to give stable dispersions was found to be 50 wt %. No stable dispersions were possible below this concentration. The average particle size of the dispersions was found to be ∼400 nm according to DLS analysis. Brij S20 as the surfactant was the most effective tenside in giving stable dispersions with OLA/DA. Replacement of DA by classical hydrophobes used for miniemulsions did not yield stable dispersions, but the addition of OA, which shows structural similarity to the DA, yielded stable dispersions in water up to 10 wt %. Stable dispersion of OLA and DA are important steps for sustainable processing from water for agricultural applications. To probe water-based processing of these dispersions, nanofiber nonwovens were prepared by electrospinning in combination with secondary aqueous dispersion of a block copolyester and a small amount of PEO. Although waterborne, these nonwovens were water stable under ambient conditions, which is another important requirement for longtime agricultural applications. Typically, for biotechnical protection against Lobesia Botrana, constant pheromone release for ∼8 weeks is required, which marks another challenge for further development of the present system. On the basis of the model release studies by isothermal thermogravimetrical analysis and previous release studies, it is suggested that further retardation of pheromone release is required and could be achieved by coating of the pheromone releasing fibers, which is the topic of upcoming contributions.

Figure 8. Comparison of the volatility of DA by thermogravimetrical analysis of free DA, DA encapsulated in OLA microparticles (1:1 by weight) and in electrospun fibers before and after water treatment by isothermal thermogravimetrical analysis for 4 h at 30 °C.

Clearly, the isothermal TGA curve for DA immobilized in OLA microparticles showed a retarded volatility in comparison with free DA. The initially faster release for DA in the OLA microparticles for the first 25 min of 2.1 wt %/h compared well with the release of free DA (1.9 wt %/h), which indicated some DA on the surface of the microparticles. For 100−200 min, a release of 0.29 wt %/h for DA in the OLA microparticles was observed, whereas for the free DA, the release rate was calculated to be 0.62 wt %/h, which indicated a retarded release of DA when it was in OLA microparticles. The strongest retardation of pheromone release was observed for electrospun fibers before water treatment, whereas after water treatment, fibers showed similar release of DA like microparticles which could support the above assumption that merging of particles occurred upon mixing of the dispersion particles of the OLA/ DA dispersion and PHA-b-MPEG dispersion. Further support for this assumption is that no indication of any microparticles originating from the OLA/DA dispersion was observed in the electrospun fibers. The consequences of this scenario would be that DA is not encapsulated in OLA microparticles in the electrospun fibers but blended in a mixture OLA and PHA-bMPEG. It is likely but still speculative that PEO does not mix well in the solid fibers with OLA and PHA-b-MPEG, which can be concluded from the porous fiber surface after removal of PEO by water extraction. This phase separation of PEO could rationalize the retarded release of DA before PEO extraction by having the fiber surface covered by a PEO layer. Nevertheless, at this state, this is speculative and cannot be proven in a sound way. However, the consequences are obvious that for an application of this system under agricultural conditions further improvement of the concept with retardation of pheromone release is required, for example, by coating of the electrospun fibers. The amount of retardation depends on the particular case, but for Lobesia Botrana

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected].

■

ACKNOWLEDGMENTS

■

REFERENCES

We thank BLE for financial support and Trifolio GmbH for the donation of pheromone.

(1) Louis, F.; Tiedemann, A. S.; Schirra, K. J. IOBC WPRS Bull. 2002, 25, 1−9. (2) Welter, S. C.; Pickel, C.; Millar, J.; Cave, F.; Steenwyk, R. A. V.; Dunley, J. Calif. Agric. 2005, 59, 16−22. (3) Shorey, H. H.; Kaae, R. S.; Gaston, L. K.; McLaughlin, J. R. Environ. Entomol. 1972, 1, 641−645. (4) Farkas, S. R.; Shorey, H. H.; Gaston, L. K. Environ. Entomol. 1974, 3, 876−877. (5) Rothschild, G. H. L. Entomol. Exp. Appl. 1974, 17, 294−302. (6) Arn, H.; Delley, B.; Baggiolini, M.; Charmillot, P. J. Entomol. Exp. Appl. 1976, 19, 139−147.

443

dx.doi.org/10.1021/bm2014679 | Biomacromolecules 2012, 13, 439−444

Biomacromolecules

Article

(7) Omi, S.; Umeki, N.; Mohri, H.; Iso, M. J. Microencapsulation 1991, 8, 465−478. (8) Minks, A. K.; Voerman, S.; Klun, J. A. Entomol. Exp. Appl. 1976, 20, 163−169. (9) Beroza, M.; Hood, C. S.; Trefrey, D.; Leonard, D. E.; Knipling, E. F.; Klassen, W.; Stevens, L. J. J. Econ. Entomol. 1974, 67, 659−664. (10) Granett, J.; Doane, C. C. J. Econ. Entomol. 1975, 68, 435−437. (11) Hellmann, C.; Greiner, A.; Wendorff, J. H. Polym. Adv. Technol. 2011, 22, 407−413. (12) Stoiljkovic, A.; Ishaque, M.; Justus, U.; Hamel, L.; Klimov, E.; Heckmann, W.; Eckhardt, B.; Wendorff, J. H.; Greiner, A. Polymer 2007, 48, 3974−3981. (13) Stoiljkovic, A.; Venkatesh, R.; Klimov, E.; Raman, V.; Wendorff, J. H.; Greiner, A. Macromolecules 2009, 42, 6147−6151. (14) Klimov, E.; Raman, V.; Venkatesh, R.; Heckmann, W.; Stark, R. Macromolecules 2010, 43, 6152−6155. (15) Sun, J.; Bubel, K.; Chen, F.; Kissel, T.; Agarwal, S.; Greiner, A. Macromol. Rapid Commun. 2010, 31, 2077−2083. (16) Beck-Broichsitter, M.; Thieme, M.; Nguyen, J.; Schmehl, T.; Gessler, T.; Seeger, W.; Agarwal, S.; Greiner, A.; Kissel, T. Macromol. Biosci. 2010, 10, 1527−1535. (17) Bognitzki, M.; Hou, H.; Ishaque, M.; Frese, T.; Hellwig, M.; Schwarte, C.; Schaper, A.; Wendorff, J. H.; Greiner, A. Adv. Mater. 2000, 12, 637−640. (18) Landfester, K. Macromol. Rapid Commun. 2001, 22, 896−936.

444

dx.doi.org/10.1021/bm2014679 | Biomacromolecules 2012, 13, 439−444