The role of phospholipid derivatives of cyclodextrins in the formation of stable lipid nanoparticles for drug delivery

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Abstract

This review article deals with physical methods for investigating the structural characteristics of inclusion complexes of supramers of phospholipid derivatives of cyclodextrins. Phospholipid derivatives of cyclodextrins are formed by attaching a phospholipid moiety to the cyclodextrin molecule. This modification imparts additional structural features to the cyclodextrin, increasing its solubility and stability in aqueous media. These new compounds can self-assemble in aqueous media into different types of supramolecular nanocomplexes. Biomedical applications are envisaged for nanoencapsulation of drug molecules in hydrophobic interchain volumes and nanocavities of amphiphilic cyclodextrins (serving as drug carriers or pharmaceutical excipients), antitumour phototherapy, gene delivery, and protection of unstable active ingredients by complexation of inclusions in nanostructured media. The focus is on the study of nanoparticle morphology, as efficient delivery systems must fulfil certain requirements. Classical physical methods cannot provide detailed information on the properties of potential structures for biomedical applications. For this purpose, the search for new non-invasive approaches is necessary.

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About the authors

E. D. Belitskaya

Shemyakin–Ovchinnikov Institute of Bioorganic Chemistry of the Russian Academy of Sciences; Moscow Institute of Physics and Technology (National Research University)

Author for correspondence.
Email: belitskayakatya@yandex.ru
Russian Federation, ul. Miklukho-Maklaya 16/10, Moscow, 117997; Institutskii per. 9, Dolgoprudny, 141701

V. A. Oleinokov

Shemyakin–Ovchinnikov Institute of Bioorganic Chemistry of the Russian Academy of Sciences; National Research Nuclear University “MEPhI”

Email: belitskayakatya@yandex.ru
Russian Federation, ul. Miklukho-Maklaya 16/10, Moscow, 117997; Kashirskoe shosse 31, Moscow, 115409

A. V. Zalygin

Shemyakin–Ovchinnikov Institute of Bioorganic Chemistry of the Russian Academy of Sciences; Lebedev Physical Institute of the Russian Academy of Sciences, Troitsk Branch

Email: belitskayakatya@yandex.ru
Russian Federation, ul. Miklukho-Maklaya 16/10, Moscow, 117997; ul. Fizicheskaya 11, Moscow, Troitsk, 108840

References

  1. Spencer D.S., Puranik A.S., Peppas N.A. // Curr. Opin. Chem. Eng. 2015. V. 7. P. 84–92. https://doi.org/10.1016/j.coche.2014.12.003
  2. Hassan S., Prakash G., Ozturk A., Saghazadeh S., Sohail M.F., Seo J., Dokmeci M., Zhang Y.S., Khademhosseini A. // Nano Today. 2017. V. 15. P. 91–106. https://doi.org/10.1016/j.nantod.2017.06.008
  3. Singh R., Lillard J.W. // Exp. Mol. Pathol. 2009. V. 86. P. 215–223. https://doi.org/10.1016/j.yexmp.2008.12.004
  4. Hu C.M.J., Fang R.H., Luk B.T., Zhang L. // Nanoscale. 2014. V. 6. P. 65–75. https://doi.org/10.1039/C3NR05444F
  5. Lakkakula J.R., Krause R.W.M. // Nanomedicine. 2014. V. 9. P. 877–894. https://doi.org/10.2217/nnm.14.41
  6. Crini G. // Chem. Rev. 2014. V. 114. P. 10940–10975. https://doi.org/10.1021/cr500081p
  7. Biwer A., Antranikian G., Heinzle E. // Appl. Microbiol. Biotechnol. 2002. V. 59. P. 609–617. https://doi.org/10.1007/s00253-002-1057-x
  8. Bonnet V., Gervaise C., Djedaïni-Pilard F., Furlan A., Sarazin C. // Drug Discov. Today. 2015. V. 20. P. 1120– 1126. https://doi.org/10.1016/j.drudis.2015.05.008
  9. Mazzaglia A., Bondì M.L., Scala A., Zito F., Barbieri G., Crea F., Vianelli G., Mineo P., Fiore T., Pellerito C., Pellerito L., Costa M.A. // Biomacromolecules. 2013. V. 14. P. 3820–3829. https://doi.org/10.1021/bm400849n
  10. Aranda C., Urbiola K., Méndez Ardoy A., García Fernández J.M., Ortiz Mellet C., de Ilarduya C.T. // Eur. J. Pharm. Biopharm. 2013. V. 85. P. 390–397. https://doi.org/10.1016/j.ejpb.2013.06.011
  11. Roux M., Sternin E., Bonnet V., Fajolles C., Djedaïni-Pilard F. // Langmuir. 2013. V. 29. P. 3677–3687. https://doi.org/10.1021/la304524a
  12. Niikura K., Matsunaga T., Suzuki T., Kobayashi S., Yamaguchi H., Orba Y., Kawaguchi A., Hasegawa H., Kajino K., Ninomiya T., Ijiro K., Sawa H. // ACS Nano. 2013. V. 7. P. 3926–3938. https://doi.org/10.1021/nn3057005
  13. Docter D., Westmeier D., Markiewicz M., Stolte S., Knauer S.K., Stauber R.H. // Chem. Soc. Rev. 2015. V. 44. P. 6094–6121. https://doi.org/10.1039/c5cs00217f
  14. Gervaise C., Bonnet V., Wattraint O., Aubry F., Sarazin C., Jaffrès P.A., Djedaïni-Pilard F. // Biochimie. 2015. V. 94. P. 66–74. https://doi.org/10.1016/j.biochi.2011.09.005
  15. Zerkoune L., Angelova A., Lesieur S. // Nanomaterials (Basel). 2014. V. 4. P. 741–765. https://doi.org/10.3390/nano4030741
  16. Auzély-Velty R., Djedaïni-Pilard F., Désert S., Perly B., Zemb T.H. // Langmur. 2000. V. 16. P. 3727–3734. https://doi.org/10.1021/la991361z
  17. Nozaki T., Maeda Y., Ito K., Kitano H. // Macromolecules. 1995. V. 28. P. 522–524. https://doi.org/10.1021/ma00106a016
  18. Kauscher U., Stuart M.C.A., Drücker P., Galla H.-J., Ravoo B.J. // Langmuir. 2013. V. 29. P. 7377–7383. https://doi.org/10.1021/la3045434
  19. Erdoğar N., Esendağlı G., Nielsen T.T., Şen M., Öner L., Bilensoy E. // Int. J. Pharm. 2016. V. 509. P. 375–390. https://doi.org/10.1016/j.ijpharm.2016.05.040
  20. Shao S., Si J., Tang J., Sui M., Shen Y. // Macromolecules. 2014. V. 47. P. 916–921. https://doi.org/10.1021/ma4025619
  21. Moutard S., Perly B., Godé P., Demailly G., Djedaïni-Pilard F. // J. Incl. Phenom. 2002. V. 44. P. 317 –322.
  22. Gèze A., Choisnard L., Putaux J.L., Wouessidjewe D. // Mater. Sci. Eng. 2009. V. 29. P. 458–462. https://doi.org/10.1016/j.msec.2008.08.027
  23. Pedersen N.R., Kristensen J.B., Bauw G., Ravoo B.J., Darcy R., Larsena K.L., Pedersen L.H. // Tetrahedron Asymmetry. 2005. V. 16. P. 615–622. https://doi.org/10.1016/j.tetasy.2004.12.009
  24. Yaméogo J.B., Gèze A., Choisnard L., Putaux J.L., Gansané A., Sirima S.B., Semdé R., Wouessidjewe D. // Eur. J. Pharm. Biopharm. 2012. V. 80. P. 508–517. https://doi.org/10.1016/j.ejpb.2011.12.007
  25. Essa S., Rabanel J.M., Hildgen P. // Int. J. Pharm. 2010. V. 388. P. 263–273. https://doi.org/10.1016/j.ijpharm.2009.12.059
  26. Bhattacharjee S. // J. Control. Release. 2016. V. 235. P. 337–351. https://doi.org/10.1016/j.jconrel.2016.06.017
  27. Lesieur S., Charon D., Lesieur P., Ringard-Lefebvre C., Muguet V., Duchêne D., Wouessidjewe D. // Chem. Phys. Lipids. 2000. V. 106. P. 127–144. https://doi.org/10.1016/S0009-3084(00)00149-3
  28. Kasselouri A., Coleman A.W., Baszkin A. // J. Colloid Interface Sci. 1996. V. 180. P. 384–397. https://doi.org/10.1006/jcis.1996.0317
  29. LoPresti C., Massignani M., Fernyhough C., Blanazs A., Ryan A.J., Madsen J., Warren N.J., Armes S.P., Lewis A.L., Chirasatitsin S., Engler A.J., Battaglia G. // ACS Nano. 2011. V. 5. P. 1775–1784. https://doi.org/10.1021/nn102455z
  30. Putaux J.L., Lancelon-Pin C., Legrand F.X., Pastrello M., Choisnard L., Gèze A., Rochas C., Wouessidjewe D. // Langmuir. 2017. V. 33. P. 7917–7928. https://doi.org/10.1021/acs.langmuir.7b01136
  31. Oliva E., Mathiron D., Rigaud S., Monflier E., Sevin E., Bricout H., Tilloy S., Gosselet F., Fenart L., Bonnet V., Pilard S., Diedaini-Pilard F. // Biomolecules. 2020. V. 10. P. 339. https://doi.org/10.3390/biom10020339
  32. Feigin L.A., Svergun D.I. // Structure Analysis by Small-Angle X-Ray and Neutron Scattering. New York: Plenum Press, 1987. V. 1. P. 14–15. https://link.springer.com/book/10.1007/978-1-4757-6624-0
  33. Auzély-Velty R., Perly B., Taché O., Zemb T., Jéhan P., Guenot P., Dalbiez J.-P., Djedaı̈ni-Pilard F. // Carbohydr. Res. 1999. V. 318. P. 82–90. https://doi.org/10.1016/S0008-6215(99)00086-5
  34. Roling O., Wendeln C., Kauscher U., Seelheim P., Galla H.-J., Ravoo B.J. // Langmuir. 2013. V. 29. P. 10174–10182. https://doi.org/10.1021/la4011218
  35. Choisnard L., Gèze A., Putaux J.L., Wong Y.S., Wouessidjewe D. // Biomacromolecules. 2006. V. 7. P. 515– 520. https://doi.org/10.1021/bm0507655
  36. Godinho B.M.D.C., Ogier J.R., Darcy R., O’Driscoll C.M., Cryan J.F. // Mol. Pharm. 2013. V. 10. P. 640–649. https://doi.org/10.1021/mp3003946
  37. Chen P., Hub J.S. // Biophys. J. 2015. V. 108. P. 2573– 2584. https://doi.org/10.1016/j.bpj.2015.03.062
  38. Vaskan I.S., Prikhodko A.T., Petoukhov M.V., Shtykova E.V., Bovin N.V., Tuzikov A.B., Oleinikov V.A., Zalygin A.V. // Colloids and Surfaces B: Biointerfaces. 2023. V. 224. P. 113183. https://doi.org/10.1016/j.colsurfb.2023.113183
  39. Zalygin A., Solovyeva D., Vaskan I., Henry S., Schaefer M., Volynsky P., Tuzikov A., Korchagina E., Ryzhov I., Nizovtsev A., Mochalov K., Efremov R., Shtykova E., Oleinikov V., Bovin N. // ChemistryOpen. 2020. V. 9. P. 641–648. https://doi.org/10.1002/open.201900276
  40. Zhou X., Liang J.F. // J. Photochem. Photobiol. A Chemistry. 2017. V. 349. P. 124–128. https://doi.org/10.1016/j.jphotochem.2017.09.032

Supplementary files

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2. Fig. 1. (a) – Functional structural diagram of α-CD (n = 6), β-CD (n = 7) and γ-CD (n = 8); (b) – geometric dimensions of cyclodextrins [6].

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3. Fig. 2. Amphiphilic cyclodextrins obtained by modifications of the macrocycle: cholesterol-cyclodextrin (a), peptide-lipidyl-cyclodextrin (b); monolauryl-cyclodextrin (c), hexanoyl-cyclodextrin (g), phospholipidyl-cyclodextrin (d); fluorinated cyclodextrin (BC6Fts: R=C6F13) (e) and octadecylperylene-cyclodextrin (g) [15].

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4. Fig. 3. Schematic representation of amphiphilic dendrimers (a) and proposed aggregate structures independently assembled from these dendrimers (b, c) [18].

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5. Fig. 4. Cryo-TEM images of multilamellar nanoparticles βCD-C10 with a total degree of substitution (TDS) = 4.3 (a–c) and βCD-C14 with TDS = 2.6 (g–e); (g) – schematic model of the transesterified derivative of βCD-C10 (TDS = 7). The grafted alkyl chains are highlighted in gray. Hydrogen atoms are omitted for clarity [30].

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6. Fig. 5. Left: Schematic view of the formation of nanoassemblers in aqueous solution from SC6OH and (Bu3Sn)4TPPSa [9] Right: Nuclear morphology of A375 melanoma cells treated with (Bu3Sn)4TPPS/SC6OH. Nomarski images (a–c, g–i, n–p); fluorescence images using Hoechst 33342 staining (d–f, j–l, r–t). Cells were treated with free (Bu3Sn)4TPPS ([(Bu3Sn)4TPPS] = 0.25 μm (a, g,w,k,n,p), nanoassemblers (Bu3Sn)4TPPS/SC6OH in a molar ratio of 1 : 5 ([(Bu3Sn)4TPPS] = 0.25 µM, [SC6OH] = 1.25 µM) (b, d, z,l, o, s) or SC6OH ([SC6OH] = 1.25 μM) (v, e, i, m, p, t) and observed nuclei after staining with the fluorescent dye Hoechst 33342 after 24, 48 and 72 h.

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7. Fig. 6. Left: Schematic representation for the preparation of folate-decorated nanocomplexes (Fol-CDplexes) from paCD T2, plasmid DNA and folic acid (FA). Right: Visualization of Balb-c mouse luciferase 24 h after intravenous administration of phosphate-buffered saline/naked DNA (a) and folate-CDplexes containing 0.5 μg FA/μg DNA (b) or 1 μg FA/μg DNA (c) [10].

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8. Fig. 7. (a) – Experimental SAXS curves measured for biotin-CMG2-DOPE nanoparticles (black dots) and simulated scattering intensities transformed back to zero angle from the distance distribution functions p(r) at different glucose concentrations: 1.0, 2.9, 3.17, and 4.23 wt %; (b) – chemical structure of biotin-CMG2-DOPE; (c) – 3D model: hydrogen atoms are highlighted in white, carbon – in blue, oxygen – in red, nitrogen – in blue, sulfur – in yellow, phosphorus – in brown; (d) – fully atomic structure of nanoparticles in cross section obtained as a result of molecular dynamics simulation; (d) – fit to the theoretical curve calculated by CRYSOL from the MD structure (red line); (f) – multiphase ab initio structure reconstruction: reconstructed electron density (color scheme: yellow corresponds to DOPE, blue – to the CMG2 spacer with biotin) and experimental data (black dots) and MONSA fit (light blue line); (g) – experimental data (black dots) and MONSA fit (light blue line); (h) – superposition of DAMMIN and MD models (transparent blue represents the DAMMIN model in phosphate buffer, and the MD CMG2 model is shown in red).

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