Helices are the most prevalent secondary structure in biomolecules and play vital roles in their activity. Chemists have been fascinated with mimicking this molecular conformation with synthetic materials. Research has now been devoted to the synthesis and characterization of helical materials, and to understand the design principles behind this molecular architecture. In parallel, work has been done to develop synthetic polymers for biological and medical applications. We now have access to materials with controlled size, molecular conformation, multivalency or functionality. As a result, synthetic polymers are being investigated in areas such as drug and gene delivery, tissue engineering, imaging and sensing, or as polymer therapeutics. Here, we provide a critical view of where these two fields, helical polymers and polymers for biological and medical applications, overlap. We have selected relevant polymer families and examples to illustrate the range of applications that can be targeted and the impact of the helical conformation on the performance. For each family of polymers, we briefly describe how they can be prepared, what helical conformations are observed and what parameters control helicity. We close this Review with an outlook of the challenges ahead, including the characterization of helicity through the process and the identification of biocompatibility.
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Yalpani, M. in Polysaccharides: Syntheses, Modifications and Structure/Property Relations 8–49 (Elsevier, 1988).
Dumitriu, S. Polysaccharides: Structural Diversity and Functional Versatility (CRC, 2004).
Pauling, L., Corey, R. B. & Branson, H. R. The structure of proteins: two hydrogen-bonded helical configurations of the polypeptide chain. Proc. Natl Acad. Sci. USA 37, 205–211 (1951).
Young, G. T. & Hardy, P. M. in Amino Acids, Peptides and Proteins Vol. 1 (ed. Young, G. T.) 112–154 (Royal Society of Chemistry, 1969).
Ruso, J. M. & Piñeiro, Á. Proteins in Solution and at Interfaces: Methods and Applications in Biotechnology and Materials Science (Wiley, 2013).
Watson, J. D. & Crick, F. H. C. Molecular structure of nucleic acids: a structure for deoxyribose nucleic acid. Nature 171, 737–738 (1953).
Franklin, R. E. & Gosling, R. G. Molecular configuration in sodium thymonucleate. Nature 171, 740–741 (1953).
Ghosh, A. & Bansal, M. A glossary of DNA structures from A to Z. Acta Crystallogr. D Biol. Crystallogr. 59, 620–626 (2003).
Nakamura, Y. in Starch: Metabolism and Structure 1–90 (Springer, 2015).
Walters, R. F. S. & DeGrado, W. F. Helix-packing motifs in membrane proteins. Proc. Natl Acad. Sci. USA 103, 13658–13663 (2006).
Barth, P. & Senes, A. Toward high-resolution computational design of the structure and function of helical membrane proteins. Nat. Struct. Mol. Biol. 23, 475–480 (2016).
Prockop, D. J. & Kivirikko, K. I. Collagens: molecular biology, diseases, and potentials for therapy. Annu. Rev. Biochem. 64, 403–434 (1995).
Karsdal, M. Biochemistry of Collagens, Laminins and Elastin: Structure, Function and Biomarkers (Elsevier Science, 2016).
Travers, A. & Muskhelishvili, G. DNA structure and function. FEBS J. 282, 2279–2295 (2015).
Nelson, D. L., Lehninger, A. L. & Cox, M. M. Lehninger Principles of Biochemistry (Rediff Books, 2017).
Hecht, S. & Huc, I. Foldamers (Wiley, 2007).
Yashima, E., Maeda, K., Iida, H., Furusho, Y. & Nagai, K. Helical polymers: synthesis, structures, and functions. Chem. Rev. 109, 6102–6211 (2009).
Nakano, T. & Okamoto, Y. in Polymer Science: A Comprehensive Reference (eds Matyjaszewski, K. & Möller, M.) 629–687 (Elsevier, 2012).
Ren, Z. & Gao, P.-X. A review of helical nanostructures: growth theories, synthesis strategies and properties. Nanoscale 6, 9366–9400 (2014).
Yashima, E. et al. Supramolecular helical systems: helical assemblies of small molecules, foldamers, and polymers with chiral amplification and their functions. Chem. Rev. 116, 13752–13990 (2016).
Green, J. J. & Elisseeff, J. H. Mimicking biological functionality with polymers for biomedical applications. Nature 540, 386–394 (2016).
Scholz, C. Polymers for Biomedicine: Synthesis, Characterization, and Applications (Wiley, 2017).
Deming, T. J. et al. Polymers at the interface with biology. Biomacromolecules 19, 3151–3162 (2018).
Yin, H. et al. Non-viral vectors for gene-based therapy. Nat. Rev. Genet. 15, 541–555 (2014).
Mitragotri, S., Burke, P. A. & Langer, R. Overcoming the challenges in administering biopharmaceuticals: formulation and delivery strategies. Nat. Rev. Drug Discov. 13, 655–672 (2014).
Stewart, M. P. et al. In vitro and ex vivo strategies for intracellular delivery. Nature 538, 183–192 (2016).
Kakkar, A., Traverso, G., Farokhzad, O. C., Weissleder, R. & Langer, R. Evolution of macromolecular complexity in drug delivery systems. Nat. Rev. Chem. 1, 0063 (2017).
Ekladious, I., Colson, Y. L. & Grinstaff, M. W. Polymer–drug conjugate therapeutics: advances, insights and prospects. Nat. Rev. Drug Discov. 18, 273–294 (2019).
Lostalé-Seijo, I. & Montenegro, J. Synthetic materials at the forefront of gene delivery. Nat. Rev. Chem. 2, 258–277 (2018).
Celiz, A. D. et al. Materials for stem cell factories of the future. Nat. Mater. 13, 570–579 (2014).
Khademhosseini, A. & Langer, R. A decade of progress in tissue engineering. Nat. Protoc. 11, 1775–1781 (2016).
Laurent, J. et al. Convergence of microengineering and cellular self-organization towards functional tissue manufacturing. Nat. Biomed. Eng. 1, 939–956 (2017).
Xia, H. et al. Tissue repair and regeneration with endogenous stem cells. Nat. Rev. Mater. 3, 174–193 (2018).
Elsabahy, M., Heo, G. S., Lim, S.-M., Sun, G. & Wooley, K. L. Polymeric nanostructures for imaging and therapy. Chem. Rev. 115, 10967–11011 (2015).
Fuchs, A. V., Gemmell, A. C. & Thurecht, K. J. Utilising polymers to understand diseases: advanced molecular imaging agents. Polym. Chem. 6, 868–880 (2015).
Yu, J., Rong, Y., Kuo, C.-T., Zhou, X.-H. & Chiu, D. T. Recent advances in the development of highly luminescent semiconducting polymer dots and nanoparticles for biological imaging and medicine. Anal. Chem. 89, 42–56 (2017).
Hu, L., Zhang, Q., Li, X. & Serpe, M. J. Stimuli-responsive polymers for sensing and actuation. Mater. Horiz. 6, 1774–1793 (2019).
Rodríguez-Hernández, J. Polymers Against Microorganisms: On the Race to Efficient Antimicrobial Materials (Springer, 2017).
Hartlieb, M., Williams, E. G. L., Kuroki, A., Perrier, S. & Locock, K. E. S. Antimicrobial polymers: mimicking amino acid functionality, sequence control and three-dimensional structure of host-defense peptides. Curr. Med. Chem. 24, 2115–2140 (2017).
Ergene, C., Yasuhara, K. & Palermo, E. F. Biomimetic antimicrobial polymers: recent advances in molecular design. Polym. Chem. 9, 2407–2427 (2018).
Deming, T. J. Synthetic polypeptides for biomedical applications. Prog. Polym. Sci. 32, 858–875 (2007).
Deng, C. et al. Functional polypeptide and hybrid materials: precision synthesis via α-amino acid N-carboxyanhydride polymerization and emerging biomedical applications. Prog. Polym. Sci. 39, 330–364 (2014).
Zagorodko, O., Arroyo-Crespo, J. J., Nebot, V. J. & Vicent, M. J. Polypeptide-based conjugates as therapeutics: opportunities and challenges. Macromol. Biosci. 17, 1600316 (2017).
Song, Z. et al. Synthetic polypeptides: from polymer design to supramolecular assembly and biomedical application. Chem. Soc. Rev. 46, 6570–6599 (2017).
Deming, T. J. in Polymer Science: A Comprehensive Reference (eds Matyjaszewski, K. & Möller, M.) 427–449 (Elsevier, 2012).
Cheng, J. & Deming, T. J. in Peptide-Based Materials (ed. Deming, T.) 1–26 (Springer, 2012).
Jiang, Z., Chen, J., Ding, J., Zhuang, X. & Chen, X. in Advances in Bioinspired and Biomedical Materials Vol. 1 (eds Ito, Y., Chen, X. & Kang, I.-K.) 149–170 (American Chemical Society, 2017).
Urnes, P. & Doty, P. in Advances in Protein Chemistry Vol. 16 (eds Anfinsen, C. B. Jr, Anson, M. L., Bailey, K. & Edsall, J. T.) 401–544 (Academic, 1962).
Katchalski, E., Sela, M., Silman, H. I. & Berger, A. in The Proteins: Composition, Structure and Function (ed. Neurath, H.) 405–602 (Academic, 1964).
Ramachandran, G. N. & Sasisekharan, V. in Advances in Protein Chemistry Vol. 23 (eds Anfinsen, C. B. Jr, Anson, M. L., Edsall, J. T. & Richards, F. M.) 283–437 (Academic, 1968).
Dill, K. A. Dominant forces in protein folding. Biochemistry 29, 7133–7155 (1990).
Chou, P. Y. & Fasman, G. D. in Advances in Enzymology and Related Areas of Molecular Biology (ed. Purich, D.) 45–148 (Wiley, 1979).
Bonduelle, C. Secondary structures of synthetic polypeptide polymers. Polym. Chem. 9, 1517–1529 (2018).
Song, Z. et al. Secondary structures in synthetic polypeptides from N-carboxyanhydrides: design, modulation, association, and material applications. Chem. Soc. Rev. 47, 7401–7425 (2018).
Chou, P. Y. & Fasman, G. D. Conformational parameters for amino acids in helical, β-sheet, and random coil regions calculated from proteins. Biochemistry 13, 211–222 (1974).
Chou, P. Y. & Fasman, G. D. Prediction of protein conformation. Biochemistry 13, 222–245 (1974).
Sasisekharan, V. Structure of poly-l-proline. II. Acta Crystallogr. 12, 897–903 (1959).
MacArthur, M. W. & Thornton, J. M. Influence of proline residues on protein conformation. J. Mol. Biol. 218, 397–412 (1991).
Adzhubei, A. A., Sternberg, M. J. E. & Makarov, A. A. Polyproline-II helix in proteins: structure and function. J. Mol. Biol. 425, 2100–2132 (2013).
Creamer, T. P. & Campbell, M. N. in Advances in Protein Chemistry Vol. 62 (ed. Rose, G. D.) 263–282 (Academic, 2002).
Shi, Z., Chen, K., Liu, Z. & Kallenbach, N. R. Conformation of the backbone in unfolded proteins. Chem. Rev. 106, 1877–1897 (2006).
Rath, A., Davidson, A. R. & Deber, C. M. The structure of “unstructured” regions in peptides and proteins: role of the polyproline II helix in protein folding and recognition. Pept. Sci. 80, 179–185 (2005).
Song, Z. et al. Enzyme-mimetic self-catalyzed polymerization of polypeptide helices. Nat. Commun. 10, 5470 (2019).
De Greef, T. F. A. et al. Supramolecular polymerization. Chem. Rev. 109, 5687–5754 (2009).
Aragonès, A. C. et al. Electrostatic catalysis of a Diels–Alder reaction. Nature 531, 88–91 (2016).
Baumgartner, R., Fu, H., Song, Z., Lin, Y. & Cheng, J. Cooperative polymerization of α-helices induced by macromolecular architecture. Nat. Chem. 9, 614–622 (2017).
Chen, C. et al. Proximity-induced cooperative polymerization in “hinged” helical polypeptides. J. Am. Chem. Soc. 141, 8680–8683 (2019).
Song, Z. et al. Synthesis of polypeptides via bioinspired polymerization of in situ purified N-carboxyanhydrides. Proc. Natl Acad. Sci. USA 116, 10658–10663 (2019).
Olander, D. S. & Holtzer, A. The stability of the polyglutamic acid alpha helix. J. Am. Chem. Soc. 90, 4549–4560 (1968).
Tomimatsu, Y., Vitello, L. & Gaffield, W. Effect of aggregation on the optical rotatory dispersion of poly(α,l-glutamic acid). Biopolymers 4, 653–662 (1966).
Saudek, V., Štokrová, Š. & Schmidt, P. Conformational study of poly(α-l-aspartic acid). Biopolymers 21, 1011–1020 (1982).
Lu, H. et al. Ionic polypeptides with unusual helical stability. Nat. Commun. 2, 206 (2011). Demonstrates that stable, water-soluble, ionic, helical poly(amino acid)s can be prepared if the helical conformation is stabilized by additional secondary interactions between the side chains, in this case, van der Waals forces.
Zhang, Y., Lu, H., Lin, Y. & Cheng, J. Water-soluble polypeptides with elongated, charged side chains adopt ultrastable helical conformations. Macromolecules 44, 6641–6644 (2011).
Engler, A. C., Lee, H. & Hammond, P. T. Highly efficient “grafting onto” a polypeptide backbone using click chemistry. Angew. Chem. Int. Ed. 48, 9334–9338 (2009).
Engler, A. C., Bonner, D. K., Buss, H. G., Cheung, E. Y. & Hammond, P. T. The synthetic tuning of clickable pH responsive cationic polypeptides and block copolypeptides. Soft Matter 7, 5627–5637 (2011).
Xiao, C. et al. Facile synthesis of glycopolypeptides by combination of ring-opening polymerization of an alkyne-substituted N-carboxyanhydride and click “glycosylation”. Macromol. Rapid Commun. 31, 991–997 (2010).
Kramer, J. R. & Deming, T. J. Preparation of multifunctional and multireactive polypeptides via methionine alkylation. Biomacromolecules 13, 1719–1723 (2012).
Kramer, J. R. & Deming, T. J. Reversible chemoselective tagging and functionalization of methionine containing peptides. Chem. Commun. 49, 5144–5146 (2013).
Kramer, J. R. et al. Reinventing cell penetrating peptides using glycosylated methionine sulfonium ion sequences. ACS Cent. Sci. 1, 83–88 (2015).
Deming, T. J. Functional modification of thioether groups in peptides, polypeptides, and proteins. Bioconjug. Chem. 28, 691–700 (2017).
Kramer, J. R. & Deming, T. J. Glycopolypeptides with a redox-triggered helix-to-coil transition. J. Am. Chem. Soc. 134, 4112–4115 (2012).
Kramer, J. R. & Deming, T. J. Multimodal switching of conformation and solubility in homocysteine derived polypeptides. J. Am. Chem. Soc. 136, 5547–5550 (2014).
Zhou, M. N. et al. N-carboxyanhydride polymerization of glycopolypeptides that activate antigen-presenting cells through dectin-1 and dectin-2. Angew. Chem. Int. Ed. 57, 3137–3142 (2018).
Kramer, J. R. & Deming, T. J. Glycopolypeptides via living polymerization of glycosylated-l-lysine N-carboxyanhydrides. J. Am. Chem. Soc. 132, 15068–15071 (2010).
Kramer, J. R., Onoa, B., Bustamante, C. & Bertozzi, C. R. Chemically tunable mucin chimeras assembled on living cells. Proc. Natl Acad. Sci. USA 112, 12574–12579 (2015). Demonstrates that helical conformations other than the α-helix can be obtained for glycosylated poly(amino acid)s, depending on the sugar conjugated and the nature of the linkage.
Brogden, K. A. Antimicrobial peptides: pore formers or metabolic inhibitors in bacteria? Nat. Rev. Microbiol. 3, 238–250 (2005).
Fjell, C. D., Hiss, J. A., Hancock, R. E. W. & Schneider, G. Designing antimicrobial peptides: form follows function. Nat. Rev. Drug Discov. 11, 37–51 (2012).
Stanzl, E. G., Trantow, B. M., Vargas, J. R. & Wender, P. A. Fifteen years of cell-penetrating, guanidinium-rich molecular transporters: basic science, research tools, and clinical applications. Acc. Chem. Res. 46, 2944–2954 (2013).
Copolovici, D. M., Langel, K., Eriste, E. & Langel, Ü. Cell-penetrating peptides: design, synthesis, and applications. ACS Nano 8, 1972–1994 (2014).
Wyrsta, M. D., Cogen, A. L. & Deming, T. J. A parallel synthetic approach for the analysis of membrane interactive copolypeptides. J. Am. Chem. Soc. 123, 12919–12920 (2001).
Koller, D. & Lohner, K. The role of spontaneous lipid curvature in the interaction of interfacially active peptides with membranes. Biochim. Biophys. Acta 1838, 2250–2259 (2014).
Xiong, M. et al. Helical antimicrobial polypeptides with radial amphiphilicity. Proc. Natl Acad. Sci. USA 112, 13155–13160 (2015).
Xiong, M. et al. Selective killing of Helicobacter pylori with pH-responsive helix–coil conformation transitionable antimicrobial polypeptides. Proc. Natl Acad. Sci. USA 114, 12675–12680 (2017).
Xiong, M. et al. Bacteria-assisted activation of antimicrobial polypeptides by a random-coil to helix transition. Angew. Chem. Int. Ed. 56, 10826–10829 (2017).
Lam, S. J. et al. Combating multidrug-resistant Gram-negative bacteria with structurally nanoengineered antimicrobial peptide polymers. Nat. Microbiol. 1, 16162 (2016).
Shirbin, S. J. et al. Architectural effects of star-shaped “structurally nanoengineered antimicrobial peptide polymers” (SNAPPs) on their biological activity. Adv. Healthc. Mater. 7, 1800627 (2018).
Engler, A. C. et al. Effects of side group functionality and molecular weight on the activity of synthetic antimicrobial polypeptides. Biomacromolecules 12, 1666–1674 (2011).
Ahmed, M. Peptides, polypeptides and peptide–polymer hybrids as nucleic acid carriers. Biomater. Sci. 5, 2188–2211 (2017).
Chen, J., Guan, X., Hu, Y., Tian, H. & Chen, X. in Polymeric Gene Delivery Systems (ed. Cheng, Y.) 85–112 (Springer, (2017).
Miyata, K., Nishiyama, N. & Kataoka, K. Rational design of smart supramolecular assemblies for gene delivery: chemical challenges in the creation of artificial viruses. Chem. Soc. Rev. 41, 2562–2574 (2012).
Mutaf, O. F., Kishimura, A., Mochida, Y., Kim, A. & Kataoka, K. Induction of secondary structure through micellization of an oppositely charged pair of homochiral block- and homopolypeptides in an aqueous medium. Macromol. Rapid Commun. 36, 1958–1964 (2015).
Perry, S. L. et al. Chirality-selected phase behaviour in ionic polypeptide complexes. Nat. Commun. 6, 6052 (2015).
Gabrielson, N. P. et al. Reactive and bioactive cationic α-helical polypeptide template for nonviral gene delivery. Angew. Chem. Int. Ed. 51, 1143–1147 (2012).
Gabrielson, N. P., Lu, H., Yin, L., Kim, K. H. & Cheng, J. A cell-penetrating helical polymer for siRNA delivery to mammalian cells. Mol. Ther. 20, 1599–1609 (2012).
Wang, H.-X. et al. Nonviral gene editing via CRISPR/Cas9 delivery by membrane-disruptive and endosomolytic helical polypeptide. Proc. Natl Acad. Sci. USA 115, 4903–4908 (2018).
Yin, L. et al. Non-viral gene delivery via membrane-penetrating, mannose-targeting supramolecular self-assembled nanocomplexes. Adv. Mater. 25, 3063–3070 (2013).
He, H. et al. Suppression of hepatic inflammation via systemic siRNA delivery by membrane-disruptive and endosomolytic helical polypeptide hybrid nanoparticles. ACS Nano 10, 1859–1870 (2016).
Liu, Y. et al. Systemic siRNA delivery to tumors by cell-penetrating α-helical polypeptide-based metastable nanoparticles. Nanoscale 10, 15339–15349 (2018).
Yin, L. et al. Light-responsive helical polypeptides capable of reducing toxicity and unpacking DNA: toward nonviral gene delivery. Angew. Chem. Int. Ed. 52, 9182–9186 (2013).
Zheng, N. et al. Manipulating the membrane penetration mechanism of helical polypeptides via aromatic modification for efficient gene delivery. Acta Biomater. 58, 146–157 (2017).
Li, F. et al. Engineering the aromaticity of cationic helical polypeptides toward “self-activated” DNA/siRNA delivery. ACS Appl. Mater. Interfaces 9, 23586–23601 (2017).
Dang, J. et al. Multivalency-assisted membrane-penetrating siRNA delivery sensitizes photothermal ablation via inhibition of tumor glycolysis metabolism. Biomaterials 223, 119463 (2019).
Pelegri-O’Day, E. M., Lin, E.-W. & Maynard, H. D. Therapeutic protein–polymer conjugates: advancing beyond PEGylation. J. Am. Chem. Soc. 136, 14323–14332 (2014).
Hou, Y. et al. Therapeutic protein PEPylation: the helix of nonfouling synthetic polypeptides minimizes antidrug antibody generation. ACS Cent. Sci. 5, 229–236 (2019).
Zhang, C. et al. From neutral to zwitterionic poly(α-amino acid) nonfouling surfaces: effects of helical conformation and anchoring orientation. Biomaterials 178, 728–737 (2018).
Cabral, H. & Kataoka, K. Progress of drug-loaded polymeric micelles into clinical studies. J. Control. Release 190, 465–476 (2014).
Cabral, H., Miyata, K., Osada, K. & Kataoka, K. Block copolymer micelles in nanomedicine applications. Chem. Rev. 118, 6844–6892 (2018).
Olsen, B. D. & Segalman, R. A. Self-assembly of rod–coil block copolymers. Mater. Sci. Eng. R. Rep. 62, 37–66 (2008).
Holowka, E. P., Pochan, D. J. & Deming, T. J. Charged polypeptide vesicles with controllable diameter. J. Am. Chem. Soc. 127, 12423–12428 (2005).
Holowka, E. P., Sun, V. Z., Kamei, D. T. & Deming, T. J. Polyarginine segments in block copolypeptides drive both vesicular assembly and intracellular delivery. Nat. Mater. 6, 52–57 (2007).
Choe, U.-J. et al. Endocytosis and intracellular trafficking properties of transferrin-conjugated block copolypeptide vesicles. Biomacromolecules 14, 1458–1464 (2013).
Schatz, C., Louguet, S., Le Meins, J. & Lecommandoux, S. Polysaccharide-block-polypeptide copolymer vesicles: towards synthetic viral capsids. Angew. Chem. Int. Ed. 48, 2572–2575 (2009).
Upadhyay, K. K. et al. The intracellular drug delivery and anti tumor activity of doxorubicin loaded poly(γ-benzyl l-glutamate)-b-hyaluronan polymersomes. Biomaterials 31, 2882–2892 (2010).
Quadir, M. A., Martin, M. & Hammond, P. T. Clickable synthetic polypeptides–routes to new highly adaptive biomaterials. Chem. Mater. 26, 461–476 (2014).
Quadir, M. A. et al. Ligand-decorated click polypeptide derived nanoparticles for targeted drug delivery applications. Nanomedicine 13, 1797–1808 (2017).
Mochida, Y. et al. Bundled assembly of helical nanostructures in polymeric micelles loaded with platinum drugs enhancing therapeutic efficiency against pancreatic tumor. ACS Nano 8, 6724–6738 (2014). Demonstrates that the helical conformation can be induced upon loading of a drug, improving not only the mechanical properties of the formed micelles but also their pharmaceutical properties.
Cabral, H. et al. Accumulation of sub-100 nm polymeric micelles in poorly permeable tumours depends on size. Nat. Nanotechnol. 6, 815–823 (2011).
Peppas, N. A. Hydrogels in Medicine and Pharmacy: Properties and Applications Vol. 3 (CRC, 2019).
Nowak, A. P. et al. Rapidly recovering hydrogel scaffolds from self-assembling diblock copolypeptide amphiphiles. Nature 417, 424–428 (2002). Demonstrates that the helical conformation has a significant impact on the mechanical properties of synthetic hydrogels, paving the way to the application of these materials for tissue engineering and drug delivery.
Breedveld, V., Nowak, A. P., Sato, J., Deming, T. J. & Pine, D. J. Rheology of block copolypeptide solutions: hydrogels with tunable properties. Macromolecules 37, 3943–3953 (2004).
Deming, T. J. Polypeptide hydrogels via a unique assembly mechanism. Soft Matter 1, 28–35 (2005).
Zhang, S., Alvarez, D. J., Sofroniew, M. V. & Deming, T. J. Design and synthesis of nonionic copolypeptide hydrogels with reversible thermoresponsive and tunable physical properties. Biomacromolecules 16, 1331–1340 (2015).
Wollenberg, A. L. et al. Injectable polypeptide hydrogels via methionine modification for neural stem cell delivery. Biomaterials 178, 527–545 (2018).
Anderson, M. A. et al. Astrocyte scar formation aids central nervous system axon regeneration. Nature 532, 195–200 (2016).
Anderson, M. A. et al. Required growth facilitators propel axon regeneration across complete spinal cord injury. Nature 561, 396–400 (2018).
Schwartz, E., Koepf, M., Kitto, H. J., Nolte, R. J. M. & Rowan, A. E. Helical poly(isocyanides): past, present and future. Polym. Chem. 2, 33–47 (2011).
Akeroyd, N., Nolte, R. J. M. & Rowan, A. E. in Isocyanide Chemistry: Applications in Synthesis and Material Science (Wiley, 2012).
Kollmar, C. & Hoffmann, R. Polyisocyanides: electronic or steric reasons for their presumed helical structure? J. Am. Chem. Soc. 112, 8230–8238 (1990).
Clericuzio, M., Alagona, G., Ghio, C. & Salvadori, P. Theoretical investigations on the structure of poly(iminomethylenes) with aliphatic side chains. Conformational studies and comparison with experimental spectroscopic data. J. Am. Chem. Soc. 119, 1059–1071 (1997).
Hase, Y. et al. Mechanism of helix induction in poly(4-carboxyphenyl isocyanide) with chiral amines and memory of the macromolecular helicity and its helical structures. J. Am. Chem. Soc. 131, 10719–10732 (2009).
Cornelissen, J. J. L. M. et al. β-Helical polymers from isocyanopeptides. Science 293, 676–680 (2001). Describes the preparation of poly(isocyanide)s with stable helicity in aqueous conditions as a result of the H-bond network formed between the peptide side chains.
Kouwer, P. H. J. et al. Responsive biomimetic networks from polyisocyanopeptide hydrogels. Nature 493, 651–655 (2013).
Das, R. K., Gocheva, V., Hammink, R., Zouani, O. F. & Rowan, A. E. Stress-stiffening-mediated stem-cell commitment switch in soft responsive hydrogels. Nat. Mater. 15, 318–325 (2016).
de Almeida, P. et al. Cytoskeletal stiffening in synthetic hydrogels. Nat. Commun. 10, 609 (2019). Reports the use of β-helical poly(isocyanide)s to form strain-stiffening gels that mimic the mechanical properties of the extracellular matrix.
op ‘t Veld, R. C. et al. Thermosensitive biomimetic polyisocyanopeptide hydrogels may facilitate wound repair. Biomaterials 181, 392–401 (2018).
Cornelissen, J. J. L. M., Fischer, M., Sommerdijk, N. A. J. M. & Nolte, R. J. M. Helical superstructures from charged poly(styrene)-poly(isocyanodipeptide) block copolymers. Science 280, 1427–1430 (1998).
Vriezema, D. M. et al. Vesicles and polymerized vesicles from thiophene-containing rod–coil block copolymers. Angew. Chem. Int. Ed. 42, 772–776 (2003). Reports the use of poly(isocyanide)s to prepare block copolymers that can afford vesicles in both organic and aqueous conditions, as a result of the unique solubility of the poly(isocyanide) block and its helical conformation.
van Oers, M. C. M., Rutjes, F. P. J. T. & van Hest, J. C. M. Cascade reactions in nanoreactors. Curr. Opin. Biotechnol. 28, 10–16 (2014).
Che, H. & van Hest, J. C. M. Adaptive polymersome nanoreactors. ChemNanoMat 5, 1092–1109 (2019).
Vriezema, D. M. et al. Positional assembly of enzymes in polymersome nanoreactors for cascade reactions. Angew. Chem. Int. Ed. 46, 7378–7382 (2007).
van Dongen, S. F. M., Nallani, M., Cornelissen, J. J. L. M., Nolte, R. J. M. & van Hest, J. C. M. A three-enzyme cascade reaction through positional assembly of enzymes in a polymersome nanoreactor. Chem. Eur. J. 15, 1107–1114 (2009).
Peters, R. J. R. W. et al. Cascade reactions in multicompartmentalized polymersomes. Angew. Chem. Int. Ed. 53, 146–150 (2014).
Peters, R. J. R. W., Louzao, I. & van Hest, J. C. M. From polymeric nanoreactors to artificial organelles. Chem. Sci. 3, 335–342 (2012).
Godoy-Gallardo, M., York-Duran, M. J. & Hosta-Rigau, L. Recent progress in micro/nanoreactors toward the creation of artificial organelles. Adv. Healthc. Mater. 7, 1700917 (2018).
van Dongen, S. F. M. et al. Cellular integration of an enzyme-loaded polymersome nanoreactor. Angew. Chem. Int. Ed. 49, 7213–7216 (2010).
Liu, J., Lam, J. W. Y. & Tang, B. Z. Acetylenic polymers: syntheses, structures, and functions. Chem. Rev. 109, 5799–5867 (2009).
Masuda, T. & Zhang, A. in Handbook of Metathesis (eds Grubbs, R. H., Wenzel, A. G., O’Leary, D. J. & Khosravi, E.) 375–390 (Wiley, 2015).
Simionescu, C. I. & Percec, V. Thermal cis–trans isomerization of cis–transoidal polyphenylacetylene. J. Polym. Sci. Polym. Chem. Ed. 18, 147–155 (1980).
Percec, V. & Rudick, J. G. Independent electrocyclization and oxidative chain cleavage along the backbone of cis-poly(phenylacetylene). Macromolecules 38, 7241–7250 (2005).
Masuda, T., Izumikawa, H., Misumi, Y. & Higashimura, T. Stereospecific polymerization of tert-butylacetylene by molybdenum catalysts. Effect of acid-catalyzed geometric isomerization. Macromolecules 29, 1167–1171 (1996).
Maeda, K. & Yashima, E. Helical polyacetylenes induced via noncovalent chiral interactions and their applications as chiral materials. Top. Curr. Chem. 375, 72 (2017).
Freire, F., Seco, J. M., Quiñoá, E. & Riguera, R. Chiral amplification and helical-sense tuning by mono- and divalent metals on dynamic helical polymers. Angew. Chem. Int. Ed. 50, 11692–11696 (2011).
Rodríguez, R., Quiñoá, E., Riguera, R. & Freire, F. Architecture of chiral poly(phenylacetylene)s: from compressed/highly dynamic to stretched/quasi-static helices. J. Am. Chem. Soc. 138, 9620–9628 (2016). A complete description of the dynamic nature of poly(acetylene)s, explored through theoretical, experimental and computational methods.
Cobos, K., Quiñoá, E., Riguera, R. & Freire, F. Chiral-to-chiral communication in polymers: a unique approach to control both helical sense and chirality at the periphery. J. Am. Chem. Soc. 140, 12239–12246 (2018).
Arias, S., Freire, F., Quiñoá, E. & Riguera, R. Nanospheres, nanotubes, toroids, and gels with controlled macroscopic chirality. Angew. Chem. Int. Ed. 53, 13720–13724 (2014).
Arias, S., Núñez-Martínez, M., Quiñoá, E., Riguera, R. & Freire, F. Simultaneous adjustment of size and helical sense of chiral nanospheres and nanotubes derived from an axially racemic poly(phenylacetylene). Small 13, 1602398 (2017).
Xu, A., Masuda, T. & Zhang, A. Stimuli-responsive polyacetylenes and dendronized poly(phenylacetylene)s. Polym. Rev. 57, 138–158 (2017).
Lv, Z., Chen, Z., Shao, K., Qing, G. & Sun, T. Stimuli-directed helical chirality inversion and bio-applications. Polymers 8, 310 (2016).
Yashima, E., Nimura, T., Matsushima, T. & Okamoto, Y. Poly((4-dihydroxyborophenyl)acetylene) as a novel probe for chirality and structural assignments of various kinds of molecules including carbohydrates and steroids by circular dichroism. J. Am. Chem. Soc. 118, 9800–9801 (1996). The demonstration that functional poly(acetylene)s can adapt their helical sense and pitch to biologically relevant metabolites, such as a glucose or steroids.
Hall, D. G. in Boronic Acids: Preparation and Applications in Organic Synthesis and Medicine 1–99 (Wiley, 2005).
Nonokawa, R. & Yashima, E. Detection and amplification of a small enantiomeric imbalance in α-amino acids by a helical poly(phenylacetylene) with crown ether pendants. J. Am. Chem. Soc. 125, 1278–1283 (2003).
Li, B. S. et al. Tuning the chain helicity and organizational morphology of an l-valine-containing polyacetylene by pH change. Nano Lett. 1, 323–328 (2001).
Arias, S., Freire, F., Calderón, M. & Bergueiro, J. Unexpected chiro-thermoresponsive behavior of helical poly(phenylacetylene)s bearing elastin-based side chains. Angew. Chem. Int. Ed. 56, 11420–11425 (2017). The demonstration that not only can thermoresponsive poly(acetylene)s be prepared but that the arrangement of the thermoresponsive moieties around a helical axis can impact the conformational changes upon heating, resulting in an unexpected increase in solubility.
Roberts, S., Dzuricky, M. & Chilkoti, A. Elastin-like polypeptides as models of intrinsically disordered proteins. FEBS Lett. 589, 2477–2486 (2015).
Bhattacharyya, J., Bellucci, J. J. & Chilkoti, A. in Biomaterials from Nature for Advanced Devices and Therapies (eds Neves, N. M. & Reis, R. L.) 106–126 (Wiley, 2016).
Freire, F., Quiñoá, E. & Riguera, R. Supramolecular assemblies from poly(phenylacetylene)s. Chem. Rev. 116, 1242–1271 (2016).
Zhao, B. & Deng, J. Emulsion polymerization of acetylenics for constructing optically active helical polymer nanoparticles. Polym. Rev. 57, 119–137 (2017).
Liang, J. & Deng, J. Chiral particles consisting of helical polylactide and helical substituted polyacetylene: preparation and synergistic effects in enantio-differentiating release. Macromolecules 51, 4003–4011 (2018).
Wang, H. et al. Chiral, thermal-responsive hydrogels containing helical hydrophilic polyacetylene: preparation and enantio-differentiating release ability. Polym. Chem. 10, 1780–1786 (2019).
Pijper, D. & Feringa, B. L. Molecular transmission: controlling the twist sense of a helical polymer with a single light-driven molecular motor. Angew. Chem. Int. Ed. 46, 3693–3696 (2007).
Lotz, B. in Synthesis, Structure and Properties of Poly(lactic acid) (eds Di Lorenzo, M. L. & Androsch, R.) 273–302 (Springer, 2018).
Thomas, S. W., Joly, G. D. & Swager, T. M. Chemical sensors based on amplifying fluorescent conjugated polymers. Chem. Rev. 107, 1339–1386 (2007).
Zhu, C., Liu, L., Yang, Q., Lv, F. & Wang, S. Water-soluble conjugated polymers for imaging, diagnosis, and therapy. Chem. Rev. 112, 4687–4735 (2012).
Liu, B. & Bazan, G. C. (eds) Conjugated Polyelectrolytes: Fundamentals and Applications (Wiley, 2013).
Kane-Maguire, L. A. P. & Wallace, G. G. Chiral conducting polymers. Chem. Soc. Rev. 39, 2545–2576 (2010).
Ho, H.-A., Najari, A. & Leclerc, M. Optical detection of DNA and proteins with cationic polythiophenes. Acc. Chem. Res. 41, 168–178 (2008).
Ho, H. et al. Colorimetric and fluorometric detection of nucleic acids using cationic polythiophene derivatives. Angew. Chem. Int. Ed. 41, 1548–1551 (2002). The demonstration that a helical conformation in conjugated polymers leads to unique spectroscopic responses that can be exploited to sense chemical mutations in DNA.
Doré, K. et al. Fluorescent polymeric transducer for the rapid, simple, and specific detection of nucleic acids at the zeptomole level. J. Am. Chem. Soc. 126, 4240–4244 (2004).
Ho, H. A. et al. Direct molecular detection of nucleic acids by fluorescence signal amplification. J. Am. Chem. Soc. 127, 12673–12676 (2005).
Nilsson, K. P. R. & Inganäs, O. Chip and solution detection of DNA hybridization using a luminescent zwitterionic polythiophene derivative. Nat. Mater. 2, 419–424 (2003).
Najari, A. et al. Reagentless ultrasensitive specific DNA array detection based on responsive polymeric biochips. Anal. Chem. 78, 7896–7899 (2006).
Fukuhara, G. & Inoue, Y. Highly selective oligosaccharide sensing by a curdlan–polythiophene hybrid. J. Am. Chem. Soc. 133, 768–770 (2011).
Nilsson, K. P. R., Rydberg, J., Baltzer, L. & Inganäs, O. Twisting macromolecular chains: self-assembly of a chiral supermolecule from nonchiral polythiophene polyanions and random-coil synthetic peptides. Proc. Natl Acad. Sci. USA 101, 11197–11202 (2004).
Sigurdson, C. J. et al. Prion strain discrimination using luminescent conjugated polymers. Nat. Methods 4, 1023–1030 (2007).
Lim, E.-K. et al. Nanomaterials for theranostics: recent advances and future challenges. Chem. Rev. 115, 327–394 (2015).
Betts, J. G. et al. in Anatomy and Physiology Ch. 3.1 (OpenStax, 2013).
P.F.-T. thanks the University of Birmingham for the John Evans Fellowship. T.L. gratefully acknowledges financial support from the Engineering and Physical Sciences Research Council (EPSRC) through a studentship from the Centre for Doctoral Training in Physical Sciences for Health (EP/L016346/1).
The authors declare no competing interests.
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- Secondary structure
The conformational arrangement (α-helix, β-pleated sheet etc.) of the backbone segments of a macromolecule, such as a polypeptide chain of a protein, without regard to the conformation of the side chains or the relationship to other segments.
The chirality of a helical, propeller or screw-shaped molecular entity.
The molecular conformation of a spiral nature, generated by regularly repeating rotations around the backbone bonds of a macromolecule.
- Helical sense
The sense of rotation around the helical axis. Viewing from either end of a molecule downwards along the helical axis, the system has P helicity (or plus) if the rotation is clockwise (or right-handed) and M helicity (or minus) if the rotation is anticlockwise (or left-handed).
- Drug delivery
The translocation of a therapeutic agent to the site of activity or infection.
- Gram-negative bacteria
Bacteria that have a thin peptidoglycan layer and an outer lipid membrane.
- Gram-positive bacteria
Bacteria that have a thick peptidoglycan layer and no outer lipid membrane.
- Gene delivery
A process by which foreign genetic material, for example, DNA or RNA, is transferred to host cells for applications such as genetic research or gene therapy. Gene delivery can result in multiple effects, including gene knockdown, i.e. the deactivation or suppression of a gene, gene knockin, i.e. the one-for-one substitution of a gene into a host’s genome, or gene knockout, i.e. the total removal or permanent deactivation of a gene.
- Gene knockdown
The deactivation or suppression of a gene.
- Helical pitch
Also known as (chain) identity period or (chain) conformational repeating unit. The distance or number of residues along the chain axis for a complete turn. This conformational unit is repeated along the chain through symmetry operations. In an α-helix formed from α-amino acids, there are 3.6 residues per ‘turn’, with a 5.4-angstroms turn.
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Leigh, T., Fernandez-Trillo, P. Helical polymers for biological and medical applications. Nat Rev Chem (2020). https://doi.org/10.1038/s41570-020-0180-5