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Keywords:

  • chitosan microspheres;
  • microfluidics;
  • drug release;
  • immobilization lipases

Chitosan, obtained by alkaline deacetylation of chitin, which is the second-most abundant polysaccharide next to cellulose, is the only base polysaccharide in nature and has outstanding properties of nontoxicity, biocompatibility, biodegradability, and low cost. Chitosan has one primary amino and two free hydroxyl groups for each C6 unit. This, in turn, makes chitosan a weak base and insoluble either in water or in organic solvents. However, it is soluble in dilute aqueous acidic solution. In the recent years, as a new type of functional materials, chitosan has great potential application in adsorption and isolation of protein, catalytic carrier, enzyme immobilization, and controlled drug release in the form of fibers, membranes, microspheres, and capsules.1

Chitosan microspheres play an important role because of their special size and structures.2 While the control of chitosan particles size and size distribution are very important to their successful biomedical applications,3 Wang et al.4 pointed out that the microspheres should be defined in a specific range, usually bellow 60 μm, and have uniform size while used as the microreactors in the field of organism catalysis and as the drug carriers. Various methods are presently available to prepare chitosan microspheres, for example, a emulsification-curing method, simple coacervation, complex coacervation, etc.5 Although the above-mentioned techniques are feasible for preparation, there are considerable drawbacks such as unstable yield, tedious procedures, non-uniform particle sizes with a wide size distribution, and lack of process repeatability. It has become imperative for the pharmaceutical industry to develop a reproducible method for generating chitosan microspheres with uniform particle size in a controlled manner, especially with size less than 60 μm. On the other hand, the control of the microspheres' structures is shown to affect the release rate and the drug dosage, which is also important to their application as drug carriers. Dambies et al.6 prepared a double-layer structure corresponding to a very compact 100-μm-thick external layer and an internal structure of small pores by preparing the chitosan gel beads in a molybdate solution under optimum conditions. Wei et al.7 prepared the chitosan microspheres with four different structures by modifying chitosan in different ways. To the best of our knowledge, there is no integrated method to prepare chitosan microspheres with different structures at present. Each structure has its own method to prepare and some of the methods are usually complicated.

Recently, microfluidic methods have been developed as the novel approaches for the controllable synthesis of monodispersed microbeads. Highly monodispersed droplets with narrow size distribution and spherical polymeric microparticles can be obtained by using microfluidic methods.8 Several research groups9 have attempted to utilize such techniques to produce chitosan microspheres, and monodispersed chitosan microspheres with controlled sizes (from 100 to 500 μm diameter) and a narrow size distribution (a variation of less than 10%) were synthesized successfully.

The requirements for the chitosan microspheres vary significantly from case to case in biomedical and biocatalysis applications. In clinical therapy, it is necessary to maintain protein drugs as a specific therapeutic serum concentration. Then the microspheres with the release profile of a minimal initial burst and a relatively well-controlled release are likely to be ideal carriers for these protein drugs. On the contrary, microspheres with a strong initial burst are suitable for treating cancer or hepatitis, which need short but intensive administration of drugs.7a In the field of biocatalysis, the enzyme should has good stability in the microspheres. Therefore, it is very important to develop an integrated and simple approach to prepare the chitosan microspheres with relatively small size (<100 μm) and controllable structures, especially in the field of drug release applications.

In this Communication, we developed a novel and simple method to prepare monodispersed chitosan microspheres with small size and controlled structures by combining the solidification methods of solvent extraction with chemical crosslinking in a capillary-embedded T-junction microfluidic device. The prepared microspheres are applied in the field of protein drug controlled release and immobilization lipases, and their properties have been studied.

In our previous work,10 the type of capillary-embedded T-junction microfluidic devices were developed to prepared monodispersed droplets. We found it could form larger shear force than other structures. So we attempted to use this type of microfluidic device to prepare monodispersed chitosan droplets and spheres with relatively small size (<100 μm). Then the chitosan droplets forming in the Teflon tube were consecutively collected with a solidification bath placed on a shaker, as shown in Figure 1a. Chitosan reacted with glutaraldehyde diffused into the droplet via the Schiff's base reaction, and the water was synchronously extracted out of the droplets by the n-octanol in the solidification bath when the droplets were shaken in the solidification bath. Therefore, the droplets gradually solidified from the surface to the inside with the residence time increasing, and monodispersed chitosan microspheres could be prepared.

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Figure 1. a) Flow chart of the experiments. b) Optical/Fluorescent micrographs of droplets and microspheres with different continuous phase flow rate at the fixed dispersed flow rate of 5.0 μL min−1. 1) Droplets, Qc = 400 μL min−1; 2) microspheres, Qc = 400 μL min−1; 3) droplets, Qc = 1550 μL min−1; 4) microspheres, Qc = 1550 μL min−1. c) The microspheres with different size by changing the solidification time at the fixed flow rates of Qc = 400 μL min−1 and Qd = 5.0 μL min−1. d) The effect of the continuous phase flow rate on the size of the microspheres with a solidification time of 35 min at a fixed dispersed flow rate of 5.0 μL min−1.

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The formed microspheres were highly monodispersed, and the microspheres' size was mainly controlled by the continuous flow rate and solidification time, as shown in Figure 1. The chitosan droplets and microspheres decreased sharply by increasing the continuous flow rate (Qc) (Figure 1b). The diameter of the microspheres changes from 130 μm to 66 μm with increasing solidification time at a fixed flow rate (Figure 1c). The average diameter of the microspheres solidified for 35 min changes from 66 μm to 25 μm with the increasing continuous phase flow rate (Figure 1d). So we successfully prepared monodispersed chitosan microspheres with relatively small size by using a novel and simple approach. The microspheres' size could be controlled from 130 μm to 25 μm simply by changing the continuous flow rate and solidification time.

The control of the microspheres' structures could also be well accomplished by regulating solidification time. In the past few decades, chitosan microspheres have always been prepared by facile chemical crosslinking using glutaraldehyde (C–G microspheres) or solvent extraction method (S–G microspheres).7, 9 Previous studies have demonstrated that C–G microspheres usually have a solid or core–shell structure without any pores on their surface,7, 9a–9e while S–G microspheres exhibit porous and core–shell structures both on the surface and on the inside.9f, 9g Thus, in an attempt to further control the structure of the chitosan microspheres, we attempted to develop a new preparation approach combining the above two methods. We can call the chitosan microspheres prepared using this novel method “CS–G microspheres”. Scheme 1 shows the mechanism of the microsphere structures generation. The chemical crosslinking reaction between glutaraldehyde and chitosan and water extraction could occur in a simultaneous way. Therefore, the size of the microspheres decreased and the structure changed with increasing solidification time. The forming process of CS–G microspheres involves three main stages with the following transformation of the microsphere structures: 1) First, the water in the droplets was extracted out, which forms pores at the microspheres' surface, allowing the porous structure to distribute no matter on the surface or in the inside. We named them “CS–G–A microspheres”. 2) With increasing solidification time, the chemical crosslinking taking place between aldehydic group and amino group can form dense structure on the surface, forming a core–shell structure with a solid shell and porous core. We named those “CS-G-B” microspheres. 3) Diffusion of glutaraldehyde molecules into the inside of the microspheres and crosslinking with the chitosan molecules results in a thicker and thicker solid shell layer, which finally leads to a complete solid structure. We named them “CS–G–C microspheres”. In other words, the structures of the microspheres can be easily controlled by simply changing the solidification time.

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Scheme 1. The mechanism of microspheres generation process. A) CS–G–A microspheres. B) CS–G–B microspheres. C) CS–G–C microspheres.

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Figure 2 shows the characterization results of the CS–G–A microspheres that were prepared under the experimental conditions of Qc = 400 μL min−1, Qd = 5.0 μL min−1, and the solidification time of 10 min. From the SEM images, the microspheres are porous both on the surface and on the inside (Figure 2a to c). The pores are mainly caused by the extraction of water from the microspheres to the solidification bath.9f Wei et al. have demonstrated the C–G microspheres show remarkable autofluorescent properties that have been attributed to the n–π* transitions of C=N bonds in the Schiff bases formed during the crosslinking reaction.7 This autofluorescent properties provide a way to characterize the thickness of the crosslinking layer of CS–G microspheres by using laser scanning confocal microscopy (LSCM), as shown in Figure 2d. The crosslinking layer attains a thickness of only a few micrometers at a solidification time of less than 10 min. A gradual decrease of the size of the surface pores and the growth of the shell thickness of these porous microspheres have been obtained by increasing the solidification time.

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Figure 2. a) Scanning electron microscopy (SEM) image of CS–G–A microspheres. b) SEM image of the surface structure of CS-G-A microspheres. c) SEM image of the inner structure of CS–G–A microspheres. d) Laser scanning confocal microscopy (LSCM) image of CS–G–A microspheres. e) SEM image of CS–G–B microspheres. f) SEM image of the surface structure of CS–G– B microspheres. g) SEM image of the inner structure of CS–G–B microspheres. h) LSCM image of CS–G–B microspheres. i) SEM image of CS–G–C microspheres. j) SEM image of the surface structure of CS–G–B microspheres. k) SEM image of the inner structure of CS–G–C microspheres with the solidification time of 35 min. l) LSCM image of CS–G–C microspheres with the solidification time of 35 min. The scale bars represent 200 μm in (a), 500 μm in (e,i), 100 μm in (d,h,l), and 50 μm in (b,c,f,g,j,k).

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As the solidification time was increased to 20 min, the chemical crosslinking taking place between the aldehydic group and the amino group formed dense structures on the surface, allowing the microspheres to form a core–shell structure with a solid shell and a porous core. CS–G–B microspheres were successfully prepared (Figure 2e–h). The thickness of the dense shell is mainly affected by the diffusion of water from microspheres into the solidification bath and the diffusion of glutaraldehyde molecules into the inside of the microspheres and crosslinking with the chitosan molecules. A gradual growth of the dense shell thickness of these CS–G–B microspheres could be obtained by increasing the solidification time.

Given a long enough solidification time (about 35 min), glutaraldehyde molecules diffused into the core of the microspheres, and the crosslinking reaction occurred everywhere in the microspheres. CS–G–C microspheres with a solid structure were formed (Figure 2i–l). The size and structure of the microspheres would no longer change with increasing solidification time, which can be seen from the comparison of Figure 2k and l with Figure S5 in the Supporting Information. The crosslinking reaction is structurally homogeneous inside the microspheres, which could enhance the mechanical stability of the microspheres.

From the above results, three typical kinds of chitosan microspheres have been successfully prepared by changing only the solidification time from 10 to 35 min, using a novel method. Compared to traditional procedures, the method presented here has enabled the preparation of monodispersed microspheres with low coefficients of variation (CV) (<5%) (Figure 2a, e, and i) and controlled size. Compared to previous microfluidic approaches, the present method gives us a simple and novel way to prepare chitosan microspheres with relatively small size and tunable structures.

The application of the prepared microspheres with different structures in the controlled release of protein-drugs and in immobilization lipases was studied. Figure 3a compares the in vitro release patterns of bovine serum albumin (BSA) from the microspheres with different structures. We found that the release ratio was below 10% in CS–G–B and CS–G–C microspheres, while it was nearly 30% in CS–G–A microspheres after 100 h. The results demonstrate that microspheres with different structures have different release profiles. CS–G–A microspheres show the highest initial burst because the macroporous outer shell and channels provide enough space for BSA to enter into the medium. CS–G–C microspheres show lowest initial burst, and the final release ratio is restricted to less than 10%, as the solid structures prevent the BSA from releasing. CS–G–B microspheres exhibit release ratios between CS–G–A and CS–G–C microspheres due to their structures of dense surfaces and porous cores. The different release profiles may meet various drug delivery requirements. CS–G–A microspheres with a large initial burst can be used for the treatment of acute diseases, while CS–G–B and CS–G–C microspheres can be used as the carriers for protein drugs that are necessary to be maintained during clinical therapy.7

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Figure 3. a) In vitro BSA release profiles measured for the different types of microspheres. b) The relationship of enzyme activity and cycle using times.

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Furthermore, for application in immobilization lipases, microspheres with compact external layers make it possible to avoid efficiently that lipases can flow away. Figure 3a shows that BSA has a lower lose rate with a longer solidification time and that the large lose rate of CS–G–B and CS–G–C microsphere is restricted to less than 10%, which shows that the lipases can exist stably in the chitosan microspheres. Figure 3b shows that the immobilization lipases in the CS–G–B and CS–G–C microspheres both exhibit about 10 times higher activity than free lipases (5 × 104 μmol g−1) and that their activities do not decrease significantly when used five times continuously; while free lipases cannot be recycled. This results illustrate that the immobilization lipase with compact external layer have both high activity and good stability. Therefore, there is great potential for chitosan microspheres with dense surfaces in the biocatalysis area. Further, this study developed a simple approach to immobilize lipases in situ.

In this Communication, a novel and simple microfluidic method was developed to prepare monodispersed chitosan microspheres with relatively small diameter and controllable structures. Highly monodispersed chitosan droplets with average diameter from 135 to 55 μm were formed by controlling the flow rate of the continuous phase via a capillary-embedded T-junction microfluidic device. The aqueous droplets formed were solidified by combining chemical crosslinking with solvent extraction. Microspheres with three typical structures, namely porous, core–shell, and solid structures, were successfully prepared by controlling the solidification time. The size of the chitosan microspheres could be controlled from 130 μm to 25 μm and it further reduced to a few micrometers with decreasing capillary diameter and increasing continuous flow rate. Furthermore, the results have shown that microspheres with different structures showed different release velocities in conrolled drug release and that in situ lipase immobilized microspheres with compact external layer had about 10 times higher activity than free lipases and showed a lower enzyme lose rate. Therefore, the monodispersed chitosan microspheres with controllable size discussed here are promising systems with better reproducibility, more repeatable release behavior, higher bioavailability, and passive targetability. The tunability of microspheres' structure enables the use of these systems to address specific requirements for use as protein drug carriers and biocatalysis.

Experimental Section

  1. Top of page
  2. Experimental Section
  3. Supporting Information
  4. Acknowledgements
  5. Supporting Information

Materials: Chitosan (0.2 g) with an average molecular weight of 180 kD (purchased from Yuhuan Ocean Biochemical Co., Ltd., Zhejiang, P. R. China) was dissolved in acetic acid (0.2 g, purchased from VAS Chemical Co., Ltd., Tianjin, P.R. China) to prepare a polymer aqueous solution (10 g). The aqueous phase was used as the dispersed phase in our experiments to form monodispersed droplets. Span80 (2 g) dissolved in n-octanol (100 g, all purchased from VAS Chemical Co., Ltd., Tianjin, P. R. China) was used as the continuous phase. Glutaraldehyde (0.06 g, purchased from VAS Chemical Co., Ltd., Tianjin, P. R. China) and Span80 (0.24 g, dissolved in n-octane (12 g) was used as the solidification bath and glutaraldehyde was used as the crosslinking reagent.

Experimental Microfluidic Device: The microfluidic device was fabricated on two 40 mm × 20 mm × 3 mm polymethyl methacrylate (PMMA) plates using micromachining technology. A Teflon tube with 0.5 mm inner diameter was inserted as the continuous phase inlet and the multiphase flow channel, while a Teflon tube with 0.05 mm inner diameter was inserted as the dispersed phase inlet (Figure S1 in the Supporting Informaiton). The microfluidic device was obtained by sealing the two PMMA plates together. Two microsyringe pumps and two gastight microsyringes were used to pump the fluids into the microfluidic device. The droplets forming in the Teflon tube were collected with a solidification bath placed on a shaker.

Preparation of Monodispersed Chitosan Microspheres: In a typical preparation experiment, the aqueous solution with 2 wt% chitosan and 2 wt% acetic acid was served as the dispersed phase, which is injected into the microchannel and separated into monodispersed droplets by the shear force of the continuous flow. An n-octanol solution with 2 wt% Span80 was used as the continuous phase. 0.5 wt% glutaraldehyde and 2 wt% Span80 added into n-octane was used as the solidification bath. The Schiff's base reaction between gluraltadehyde and chitosan and the extraction of water by n-octanol were employed to solidify the droplets in the solidification bath. The microspheres with different solidification time were obtained by controlling the time droplets shaken in the solidification bath. Finally, the spheres were washed with n-octane and dried by freeze-drying.

In situ Preparation of BSA-Loaded and Lipases-Loaded Chitosan Microspheres: BSA is used as the protein drug model in our experiment. The continuous phase and the solidification bath were as same as these described above. Besides the component of the dispersed phase, 0.2 wt% BSA was added and, at the same time, the quantity of acetic acid was reduced from 2 wt% to 0.33 wt% in order to avoid the BSA losing its activity. The dispersed phase should be used after centrifuging. The formation of the microspheres is identical to the process presented above. The lipase-coated in situ chitosan microspheres can be obtained by putting the same quantity of lipase into the dispersed phase instead of the BSA.

Characterization: Droplets and microspheres were observed with an optical microscope (Olympus) and an on-line CCD (Pixlink). More detailed structures were observed using scanning electron microscopy (SEM, FEI XL30). Fluorescence was observed using laser sanning confocal microscopy (LSM710, Zeiss). The BSA/lipases-loaded spheres were characterized by desorption of BSA and the enzyme activity of lipases. The BSA release experiment was performed by putting the BSA-coated in situ chitosan microspheres into the phosphoric acid buffer solution (pH = 7.0, 0.05 M), shaking from 1.0 h to 100 h, and then determining the BSA concentration of the solution using UV–vis spectrophotometer.7 The standard enzyme activity experiment in aqueous media is the hydrolyzation reaction of acetic acid glyceride in 30 min.

Supporting Information

  1. Top of page
  2. Experimental Section
  3. Supporting Information
  4. Acknowledgements
  5. Supporting Information

Supporting Information is available from the Wiley Online Library or from the author.

Acknowledgements

  1. Top of page
  2. Experimental Section
  3. Supporting Information
  4. Acknowledgements
  5. Supporting Information

This work was supported by the National Natural Science Foundation of China (21036002, 21136006, 20806042) and A Foundation for the Author of National Excellent Doctoral Dissertation of PR China (FANEDD 201053).

Supporting Information

  1. Top of page
  2. Experimental Section
  3. Supporting Information
  4. Acknowledgements
  5. Supporting Information

Detailed facts of importance to specialist readers are published as ”Supporting Information”. Such documents are peer-reviewed, but not copy-edited or typeset. They are made available as submitted by the authors.

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