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Final Report Summary - SAMUL-NANO-HEP (Self-Assembling Multivalent Biodegradable Ligands for Nanoscale Heparin Binding)

Our main goal was to develop nanoscale strategies to understand how polyanionic heparin, used during major surgery as an anti-coagulant, can be effectively bound, and removed once surgery is complete. We therefore aimed to develop heparin binders with potential clinical relevance.

We synthesised a novel family of peptide-derived molecules combining hydrophobic modification with cationic amino acid terminal groups.1 The hydrophobic groups are responsible for self-assembly, and the resulting self-assembled multivalent (SAMul) amino acids bind anionic heparin (see Figure). The units are connected with linkers that can degrade, disassembling the nano-structure and turning off binding. Specifically, we employed a lysine group as a heparin binder to create a rod-like ‘first generation’ (G1) binder. This lysine was then used as a branching group to allow the synthesis of more dendritic cone-like ‘second generation’ (G2) ligands. The units were connected through a central aspartic acid linker. We also modified chirality as well as the hydrophobic unit chain length to yield a family of twin-tailed surfactants. Some simple ligands were also prepared for use as reference molecules.

We monitored the ability of these compounds to self-assemble using a Nile Red assay, determining the critical aggregation concentrations (CACs).2 CAC values for the twin-tailed G1 systems were significantly larger than those for single-tailed systems developed previously in the group. Dynamic light scattering (DLS) was performed to gain insight into sizes and charges densities of the resulting aggregates. Rod-like G1 systems gave rise to much larger assembled structures than cone-like G2 systems (see Figure). We propose that the G2 ligands form small well-defined spherical micellar assemblies, while G1 systems form less structurally-defined larger lamellar self-assembled structures. Zeta-potential measurements indicated the larger G1 assemblies were significantly more charge dense. We also explored the impact of environment (e.g. salt levels) on self-assembly, as such factors are important in vivo. Transmission electron microscopy (TEM) was supportive of DLS observations – larger vesicular-type objects were observed for G1 – in some cases, these appeared to be multi-lamellar, with several layers being distinguishable within the nanoscale objects. For G2, small spherical micelles were dominant, some of which assembled into larger aggregates.

We then determined the relative heparin binding affinities using an assay, monitoring displacement of Mallard Blue dye from its complex with heparin.3 The G2 systems, which form small micellar assemblies, were much more effective heparin binders than the G1 systems. To understand this in more detail we performed multiscale modelling in collaboration with Prof Sabrina Pricl (University of Trieste). Interestingly, although the small micellar assemblies formed by the larger G2 monomers exhibit better heparin binding, their performance was significantly perturbed in human serum compared with G1.4 We suggest that the known susceptibility of spherical micelles to disruption in serum and the greater stability of lamellar vesicular assemblies underpins this performance difference. Importantly, this would suggest that structures which assemble into vesicles are optimal for this application in vivo. As such, our structure-activity relationship study provided important information to optimise these self-assembling nanostructures for clinically-relevant heparin rescue.

We also compared the heparin binding of the ligands with the ability to bind DNA using an ethidium bromide displacement assay.5 Interesting results were obtained with enantiomeric pairs of self-assemblies; the data showed the unlike when binding heparin, the enantiomeric systems bound DNA with very different charge efficiencies, suggesting that the DNA binding interface may have greater chiral ‘definition’ than that of heparin – as might be expected from the repeating helical structure.6

As part of this project, we also developed a high-throughput Mallard Blue assay to test a variety of mixed co-assembled binders. This rapidly allowed us to identify any systems which were outperforming, or underperforming, and hence single them out for further study and fuller titration analysis. For some mixtures we could observed that the effects of mixing were, perhaps surprisingly, non-linear, with one of the binders tending to dominate the process, and the other to disrupt it. This high-throughput approach allowed efficient exploration of chemical space and optimisation of co-assembled binding systems.

We finally probed the degradation of our self-assembled nanostructures under ambient physiological conditions of pH using mass spectrometry. In all cases, degradation of the molecules was observed over a 24 hour time period – this is clinically useful as any excess heparin binder will be simply degraded in vivo into non-self-assembling, non-multivalent, non-toxic and non-bioactive fragments. This approach could therefore transform treatment approaches for heparin rescue, which are currently very conservative owing to the side effects of the clinical agent, protamine.


In summary, the work developed here allowed us to gain a much more detailed and rational understanding of the key features required for heparin binding, such as charge/shape/size, and also explore how it may be possible to discriminate between heparin and other negatively charged nanoscale targets. These results are of significant to chemists working in nanoscale supramolecular chemistry and have potential impact in clinical surgical applications. Optimised heparin binders developed as a result of knowledge from this project are being further developed for potential clinical translation.

1. A. Barnard and D. K. Smith, Angew. Chem. Int. Ed. 2012, 51, 6572-6581.
2. M. C. A. Stuart, J. C. van de Pas and J. B. F. N. Engberts, J. Phys. Org. Chem. 2005, 18, 929-934.
3. (a) S. M. Bromfield, A. Barnard, P. Posocco, M. Fermeglia, S. Pricl and D. K. Smith, J. Am. Chem. Soc., 2013, 135, 2911-2914 (b) S. M. Bromfield, P. Posocco, M. Fermeglia, S. Pricl, J. Rodríguez-López and D. K. Smith, Chem. Commun., 2013, 49, 4830-4832.
4. (a) O. Freund, J. Amédee, D. Roux and R. Laversanne, Life Sci., 2000, 67, 411-419 (b) J. Lu, S. C. Owen and M. S. Shoichet, Macromolecules, 2011, 44, 6002-6008.
5. (a) B. F. Cain, B. C. Baguley and W. A. Denny, J. Med. Chem. 1978, 21, 658-668. (b) H. Gershon, R. Ghirlando, S. B. Guttman and A. Minsky, Biochemistry 1993, 32, 7143-7151
6. S. M. Bromfield and D. K. Smith, J. Am. Chem. Soc., 2015, 137, 10056–10059.

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