BioChip Journal

, Volume 7, Issue 3, pp 210–217 | Cite as

High-throughput nanoscale lipid vesicle synthesis in a semicircular contraction-expansion array microchannel

  • Jisun Lee
  • Myung Gwon Lee
  • Cheulhee Jung
  • Youn-Hee Park
  • Chaeyeon Song
  • Myung Chul Choi
  • Hyun Gyu Park
  • Je-Kyun Park
Original Article

Abstract

Towards potential applications in the field of nanomedicine, a new high-throughput synthesis method of lipid vesicles with tunable size as well as enhanced monodispersity is demonstrated using a semicircular contraction-expansion array (CEA) microchannel. Lipid vesicles are generated in the CEA microchannel by injecting lipids in isopropyl alcohol as a sample flow and phosphate buffered saline as a buffer flow, leading to spontaneous formation of lipid vesicles. In the CEA microchannel, Dean vortices cause three-dimensional (3D) lamination by continuously splitting and redirecting fluid streams, resulting in enhancement of fluid mixing. When considered only 3D laminating effect, it showed the best mixing efficiency in the range of flow rates of 12–15 mL/h. However, shear force effect also gives a strong influence on the formation of lipid vesicles, leading to the smallest size and uniform size distribution of lipid vesicles at a total flow rate of 18 mL/h. Consequently, from the interplay between high shear stress and 3D laminating effect, the lipid vesicles were generated with monodispersity and high throughput. The formation of lipid vesicles can be controlled with a total flow rate and a flow rate ratio between the sample and buffer fluids. The throughput of the lipid generation in the CEA microchannel was 10 times higher than previous works. In addition, the generated lipid vesicle populations were confirmed using a cryogenic transmission electron microscopy (cryo-TEM) technique.

Keywords

Contraction-expansion array microchannel High-throughput synthesis Lipid vesicles Microfluidics Monodispersity 

Preview

Unable to display preview. Download preview PDF.

Unable to display preview. Download preview PDF.

References

  1. 1.
    Jahn, A. et al. Microfluidic directed formation of liposomes of controlled size. Langmuir 23, 6289–6293 (2007).CrossRefGoogle Scholar
  2. 2.
    Hong, J.S. et al. Liposome-templated supramolecular assembly of responsive alginate nanogels. Langmuir 24, 4092–4096 (2008).CrossRefGoogle Scholar
  3. 3.
    Jahn, A., Vreeland, W.N., Gaitan, M. & Locascio, L.E. Controlled vesicle self-assembly in microfluidic channels with hydrodynamic focusing. J. Am. Chem. Soc. 126, 2674–2675 (2004).CrossRefGoogle Scholar
  4. 4.
    Huang, X.M. et al. Ultrasound-enhanced microfluidic synthesis of liposomes. Anticancer Res. 30, 463–466 (2010).Google Scholar
  5. 5.
    Gullotti, E. & Yeo, Y. Extracellularly activated nanocarriers: a new paradigm of tumor targeted drug delivery. Mol. Pharmaceut. 6, 1041–1051 (2009).CrossRefGoogle Scholar
  6. 6.
    Ishida, T., Harashima, H. & Kiwada, H. Liposome clearance. Bioscience. Rep. 22, 197–224 (2002).CrossRefGoogle Scholar
  7. 7.
    Xu, Q. et al. Preparation of monodisperse biodegradable polymer microparticles using a microfluidic flow-focusing device for controlled drug delivery. Small 5, 1575–1581 (2009).CrossRefGoogle Scholar
  8. 8.
    Traikia, M. et al. Formation of unilamellar vesicles by repetitive freeze-thaw cycles: characterization by electron microscopy and P-31-nuclear magnetic resonance. Eur. Biophys. J. Biophy. 29, 184–195 (2000).CrossRefGoogle Scholar
  9. 9.
    Tan, Y.C., Hettiarachchi, K., Siu, M. & Pan, Y.P. Controlled microfluidic encapsulation of cells, proteins, and microbeads in lipid vesicles. J. Am. Chem. Soc. 128, 5656–5658 (2006).CrossRefGoogle Scholar
  10. 10.
    Batzri, S. & Korn, E.D. Single bilayer liposomes prepared without sonication. Biochim. Biophys. Acta 298, 1015–1019 (1973).CrossRefGoogle Scholar
  11. 11.
    Maulucci, G. et al. Particle size distribution in DMPC vesicles solutions undergoing different sonication times. Biophys. J. 88, 3545–3550 (2005).CrossRefGoogle Scholar
  12. 12.
    Jahn, A. et al. Microfluidic mixing and the formation of nanoscale lipid vesicles. ACS Nano 4, 2077–2087 (2010).CrossRefGoogle Scholar
  13. 13.
    Valencia, P.M. et al. Single-step assembly of homogenous lipid — polymeric and lipid-quantum dot nanoparticles enabled by microfluidic rapid mixing. ACS Nano 4, 1671–1679 (2010).CrossRefGoogle Scholar
  14. 14.
    Ramachandran, S., Quist, A.P., Kumar, S. & Lal, R. Cisplatin nanoliposomes for cancer therapy: AFM and fluorescence Imaging of cisplatin encapsulation, stability, cellular uptake, and toxicity. Langmuir 22, 8156–8162 (2006).CrossRefGoogle Scholar
  15. 15.
    Abraham, S.A. et al. The liposomal formulation of doxorubicin. Method Enzymol. 391, 71–97 (2005).CrossRefGoogle Scholar
  16. 16.
    Gulsen, D., Li, C.C. & Chauhan, A. Dispersion of DMPC liposomes in contact lenses for ophthalmic drug delivery. Curr. Eye Res. 30, 1071–1080 (2005).CrossRefGoogle Scholar
  17. 17.
    Andresen, T.L., Jensen, S.S. & Jorgensen, K. Advanced strategies in liposomal cancer therapy: problems and prospects of active and tumor specific drug release. Prog. Lipid Res. 44, 68–97 (2005).CrossRefGoogle Scholar
  18. 18.
    Crosasso, P. et al. Preparation, characterization and properties of sterically stabilized paclitaxel-containing liposomes. J. Control. Release 63, 19–30 (2000).CrossRefGoogle Scholar
  19. 19.
    Sadava, D., Coleman, A. & Kane, S.E. Liposomal daunorubicin overcomes drug resistance in human breast, ovarian and lung carcinoma cells. J. Liposome Res. 12, 301–309 (2002).CrossRefGoogle Scholar
  20. 20.
    Pavelic, Z. et al. Development and in vitro evaluation of a liposomal vaginal delivery system for acyclovir. J. Control. Release 106, 34–43 (2005).CrossRefGoogle Scholar
  21. 21.
    Boonyasit, Y. et al. Passive micromixer integration with a microfluidic chip for calcium assay based on the arsenazo III method. BioChip J. 5, 1–7 (2011).CrossRefGoogle Scholar
  22. 22.
    Karnik, R. et al. Microfluidic platform for controlled synthesis of polymeric nanoparticles. Nano Lett. 8, 2906–2912 (2008).CrossRefGoogle Scholar
  23. 23.
    Stroock, A.D. et al. Chaotic mixer for microchannels. Science 295, 647–651 (2002).CrossRefGoogle Scholar
  24. 24.
    Lin, Y.C., Chung, Y.C. & Wu, C.Y. Mixing enhancement of the passive microfluidic mixer with J-shaped baffles in the tee channel. Biomed. Microdevices 9, 215–221 (2007).CrossRefGoogle Scholar
  25. 25.
    Hong, C.C., Choi, J.W. & Ahn, C.H. A novel inplane passive microfluidic mixer with modified Tesla structures. Lab Chip 4, 109–113 (2004).CrossRefGoogle Scholar
  26. 26.
    Lee, M.G., Choi, S. & Park, J.-K. Rapid laminating mixer using a contraction-expansion array microchannel. Appl. Phys. Lett. 95, 051902 (2009).CrossRefGoogle Scholar
  27. 27.
    Sudarsan, A.P. & Ugaz, V.M. Fluid mixing in planar spiral microchannels. Lab Chip 6, 74–82 (2006).CrossRefGoogle Scholar
  28. 28.
    Howell, P.B., Mott, D.R., Golden, J.P. & Ligler, F.S. Design and evaluation of a Dean vortex-based micromixer. Lab Chip 4, 663–669 (2004).CrossRefGoogle Scholar
  29. 29.
    Lee, M.G., Choi, S. & Park, J.K. Rapid multivortex mixing in an alternately formed contraction-expansion array microchannel. Biomed. Microdevices 12, 1019–1026 (2010).CrossRefGoogle Scholar
  30. 30.
    Zhang, H.W. et al. Assembly of plasmid DNA into liposomes after condensation by cationic lipid in anionic detergent solution. Biotechnol. Lett. 27, 1701–1705 (2005).CrossRefGoogle Scholar

Copyright information

© The Korean BioChip Society and Springer-Verlag Berlin Heidelberg 2013

Authors and Affiliations

  • Jisun Lee
    • 1
  • Myung Gwon Lee
    • 1
  • Cheulhee Jung
    • 2
  • Youn-Hee Park
    • 1
  • Chaeyeon Song
    • 1
  • Myung Chul Choi
    • 1
  • Hyun Gyu Park
    • 2
  • Je-Kyun Park
    • 1
  1. 1.Department of Bio and Brain EngineeringKAISTDaejeonKorea
  2. 2.Department of Chemical and Biomolecular EngineeringKAISTDaejeonKorea

Personalised recommendations