bind to the surface. Surfactants that form stable complexes with proteins (e.g. poloxamers) and protect them by colloidal stabilization are not showing accelerated adsorption in the presence of protein.

## 6.5 Perspectives on molecular surface adsorption of mAbs and surfactants

In order to guide formulation optimization, it will be important to establish the underlying molecular protection mechanism in existing formulations. Considering the studied formulations, our approach will be the following:

* Determine apparent $k_{on}$ rates for mAbs and surfactants from kinetic SPRi data

* Use $k_{on}$ rates to predict if mAb or surfactant will bind first at the hydrophobic surface

* Demonstrate competitive binding of mAb or surfactant in formulation conditions

* Analyse stability of mAb under dehydration stress (air interface) with and without surfactants

* Establish molecular protection mechanism from the above results

We will analyse a series of mAb formulations with different types of immunoglobulins and surfactants on selected surface chemistries. The protocols for the kinetic analyses by SPRi are established and will be used to determine apparent $k_{on}$ rates for the adsorption of the different molecules. The adsorption behaviour in formulation conditions can then be predicted and quantitatively verified. For this, we will develop an “ELISA-type” protocol using a secondary Ab or proteinA/G-based conjugate that allows to identify and quantify adsorbed mAb when a surface is incubated with the complete formulation, the formulation without mAb and the formulation without surfactant. This can be done using the SPRi signal of the binding secondary Ab or proteinA/G, or in a 96-well microplate protocol with an enzymatic readout.

The protection of the mAbs from dehydration stress at the air interface will also be analysed. As seen in chapter 5.5 (Insulin aggregation at a dynamic triple interface) the dehydration provoked by the movement of a sweeping solution meniscus over a hydrophobic surface with adsorbed proteins triggers their aggregation at the triple line. We will thus analyse the surface-adsorption state of mAbs in the presence or absence of surfactants, when they are transiently
exposed to air. SPRi allows to follow the adsorbed protein amount in these conditions and can therefore be used to study potential desorption. Conformational transitions induced by a transient exposure to air can be measured by infrared spectroscopy (ATR-FTIR) or with fluorescent conformational markers (e.g. ProteoStat). Obtaining images of adsorbed protein and surfactant layers in different conditions and on different surface chemistries is another useful complementary approach. Surface-enhanced ellipsometric contrast (SEEC) imaging is well suited for this. It is based on engineered glass coatings (commercialized by Nanolane), which allow to precisely control the surface reflection properties by multiple reflections and interferences such that the polarisation of the reflected light is not changed (Figure 6-8)¹⁶⁴. In this way the contrast of nanometric objects deposited on them can be optimized. This is highly relevant for thin layers of translucent material that has very low optical contrast.

<figure><img src="image_54.png" /><figcaption>Figure 6-8: Principle of SEEC microscopy. Left A substrate (1) is covered by optical layers (2) providing a completely dark background (no change in the polarisation of the reflected light). Right When a small object binds to the surface (4), its presence is detected with nanometric axial sensitivity. Figure credits to Nanolane (http://www.nano-lane.com/seec-technology/principles/)</figcaption></figure>

This interference microscopy technique can be used in liquid and in dry conditions (air) and is ideally suited to study protein adsorption at functionalized surfaces¹⁶⁵,¹⁶⁶ as its axial (z) resolution is below 1 nm (Figure 6-9).

<figure><img src="image_55.png" /><figcaption>Figure 6-9: Wet-SEEC image (left) and Wet-SEEC topographic reconstruction (right) of calibrated resin steps, with thicknesses of 10, 18, and 40 nm. Scale bar, 10 µm. Figure adapted from¹⁶⁶.</figcaption></figure>
In order to understand the molecular mechanisms underlying protein and surfactant surface adsorption it is important to obtain images of surface coverage and to measure surface-adsorbed molecular layer dimensions in real time. A detailed and more comprehensive picture of what happens to proteins and surfactants at liquid-solid and liquid-air interfaces can thus be obtained. Of special interest will be the SEECS analysis of the impact of dehydration. Measuring the variation of adsorbed layer thicknesses in response to dehydration can inform on the protection mechanism of surfactants which might help retaining water around adsorbed proteins (Figure 6-10). In this way, intermolecular protein aggregation triggered by the small distance between surface-adsorbed molecules and conformational changes due to variations in their hydration shell could be prevented.

<figure><img src="image_56.png" /><figcaption>Figure 6-10: Scheme representing surface-adsorbed molecules (blue) which, when exposed to air, can be dehydrated (top). In the presence of surfactants (red) a protective water layer is retained (bottom).</figcaption></figure>

The picture of how proteins and other molecules behave at interfaces is far from being completed. The infinite complexity of proteins and the simultaneous occurrence of different forces and phenomena at interfaces make this subject a rich and rewarding research area, abounding our fundamental knowledge in life sciences and applied research alike.
# 7 References

1. Hartl, F. U., Bracher, A. & Hayer-Hartl, M. Molecular chaperones in protein folding and proteostasis. *Nature* **475**, 324–332 (2011).

2. Saibil, H. Chaperone machines for protein folding, unfolding and disaggregation. *Nat Rev Mol Cell Biol* **14**, 630–642 (2013).

3. Anfinsen, C. B., Haber, E., Sela, M. & White, F. H. The kinetics of formation of native ribonuclease during oxidation of the reduced polypeptide chain. *Proc. Natl. Acad. Sci. U. S. A.* **47**, 1309–1314 (1961).

4. Jahn, T. R. & Radford, S. E. The Yin and Yang of protein folding. *FEBS J* **272**, 5962–5970 (2005).

5. Dragoš, A., Kovács, Á. T. & Claessen, D. The Role of Functional Amyloids in Multicellular Growth and Development of Gram-Positive Bacteria. *Biomolecules* **7**, 60 (2017).

6. Turoverov, K. K., Kuznetsova, I. M. & Uversky, V. N. The protein kingdom extended: ordered and intrinsically disordered proteins, their folding, supramolecular complex formation, and aggregation. *Prog Biophys Mol Biol* **102**, 73–84 (2010).

7. Fischer, E. Einfluss der Configuration auf die Wirkung der Enzyme. *Berichte der Dtsch. Chem. Gesellschaft* **27**, 2985–2993 (1894).

8. He, B. et al. Predicting intrinsic disorder in proteins: an overview. *Cell Res* **19**, 929–949 (2009).

9. Mollica, L. et al. Binding Mechanisms of Intrinsically Disordered Proteins: Theory, Simulation, and Experiment. *Front Mol Biosci* **3**, 52 (2016).

10. Zhou, H. X. Intrinsic disorder: signaling via highly specific but short-lived association. *Trends Biochem Sci* **37**, 43–48 (2012).

11. Novo, M., Freire, S. & Al-Soufi, W. Critical aggregation concentration for the formation of early Amyloid-β (1–42) oligomers. *Sci. Rep.* **8**, 1783 (2018).

12. Banerjee, S. et al. A novel pathway for amyloids self-assembly in aggregates at nanomolar concentration mediated by the interaction with surfaces. *Sci. Rep.* **7**, (2017).

13. Baldwin, A. J. et al. Metastability of native proteins and the phenomenon of amyloid formation. *J. Am. Chem. Soc.* **133**, 14160–14163 (2011).

14. Auer, S., Meersman, F., Dobson, C. M. & Vendruscolo, M. A generic mechanism of emergence of amyloid protofilaments from disordered oligomeric aggregates. *PLoS Comput. Biol.* **4**, (2008).

15. Eanes, E. D. & Glenner, G. G. X-ray diffraction studies on amyloid filaments. *J. Histochem. Cytochem.* **16**, 673–677 (1968).

16. Knowles, T. P. & Buehler, M. J. Nanomechanics of functional and pathological amyloid materials. *Nat Nanotechnol* **6**, 469–479 (2011).

17. Serpell, L. Amyloid structure. *Essays Biochem.* **56**, 1–10 (2014).

18. Jimenez, J. L. et al. The protofilament structure of insulin amyloid fibrils. *Proc Natl Acad Sci USA* **99**, 9196–9201 (2002).

19. Glover, J. R. et al. Self-Seeded Fibers Formed by Sup35, the Protein Determinant of PSI+, a Heritable Prion-like Factor of S. cerevisiae. *Cell* **89**, 811–819 (1997).

20. Knowles, T. P. J. et al. Observation of spatial propagation of amyloid assembly from single nuclei. *Proc. Natl. Acad. Sci.* **108**, 14746–14751 (2011).
21. Fodera, V., Librizzi, F., Groenning, M., van de Weert, M. & Leone, M. Secondary nucleation and accessible surface in insulin amyloid fibril formation. *J Phys Chem B* **112**, 3853–3858 (2008).

22. Agrawal, S. G. & Paterson, A. H. J. Secondary Nucleation: Mechanisms and Models. *Chemical Engineering Communications* **202**, 698–706 (2015).

23. Forman-Kay, J. D. & Mittag, T. From sequence and forces to structure, function, and evolution of intrinsically disordered proteins. *Structure* **21**, 1492–1499 (2013).

24. Tyedmers, J., Mogk, A. & Bukau, B. Cellular strategies for controlling protein aggregation. *Nat Rev Mol Cell Biol* **11**, 777–788 (2010).

25. Scior, A., Juenemann, K. & Kirstein, J. Cellular strategies to cope with protein aggregation. *Essays Biochem.* **60**, 153–161 (2016).

26. Nakatogawa, H., Suzuki, K., Kamada, Y. & Ohsumi, Y. Dynamics and diversity in autophagy mechanisms: lessons from yeast. *Nat. Rev. Mol. Cell Biol.* **10**, 458–467 (2009).

27. Rapoport, T. A., Li, L. & Park, E. Structural and Mechanistic Insights into Protein Translocation. *Annu. Rev. Cell Dev. Biol.* **33**, annurev-cellbio-100616-060439 (2017).

28. Xu, J., Wang, J., Wimo, A. & Qiu, C. The economic burden of mental disorders in China, 2005-2013: implications for health policy. *BMC Psychiatry* **16**, 137 (2016).

29. Knowles, T. P. J., Vendruscolo, M. & Dobson, C. M. The amyloid state and its association with protein misfolding diseases. *Nat. Rev. Mol. Cell Biol.* **15**, 384–396 (2014).

30. Stefani, M. & Dobson, C. M. Protein aggregation and aggregate toxicity: new insights into protein folding, misfolding diseases and biological evolution. *J Mol Med* **81**, 678–699 (2003).

31. Baglioni, S. et al. Prefibrillar Amyloid Aggregates Could Be Generic Toxins in Higher Organisms. *J. Neurosci.* **26**, 8160–8167 (2006).

32. Olzscha, H. et al. Amyloid-like aggregates sequester numerous metastable proteins with essential cellular functions. *Cell* **144**, 67–78 (2011).

33. Bulawa, C. E. et al. Tafamidis, a potent and selective transthyretin kinetic stabilizer that inhibits the amyloid cascade. *Proc. Natl. Acad. Sci.* **109**, 9629–9634 (2012).

34. Ecker, D. M., Jones, S. D. & Levine, H. L. The therapeutic monoclonal antibody market. *MAbs* **7**, 9–14 (2015).

35. Strohl, W. R. Current progress in innovative engineered antibodies. *Protein and Cell* **9**, 86–120 (2018).

36. Jozala, A. F. et al. Biopharmaceuticals from microorganisms: from production to purification. *Brazilian J. Microbiol.* **47**, 51–63 (2016).

37. Karst, D. J., Steinebach, F. & Morbidelli, M. Continuous integrated manufacturing of therapeutic proteins. *Current Opinion in Biotechnology* **53**, 76–84 (2018).

38. Kolhe, P., Amend, E. & Singh, S. K. Impact of freezing on pH of buffered solutions and consequences for monoclonal antibody aggregation. *Biotechnol Prog* **26**, 727–733 (2010).

39. Privalov, P. L. Cold denaturation of proteins. *Crit Rev Biochem Mol Biol* **25**, 281–305 (1990).

40. Padala, C., Jameel, F., Rathore, N., Gupta, K. & Sethuraman, A. Impact of Uncontrolled vs Controlled Rate Freeze-Thaw Technologies on Process Performance and Product Quality. *PDA J Pharm Sci Technol* **64**, 290–298 (2010).
41. Rathore, N. & Rajan, R. S. Current Perspectives on Stability of Protein Drug Products during Formulation, Fill and Finish Operations. *Biotechnol Prog* **24**, 504–514 (2008).

42. Mahler, H.-C., Printz, M., Kopf, R., Schuller, R. & Müller, R. Behaviour of Polysorbate 20 During Dialysis, Concentration and Filtration Using Membrane Separation Techniques. *J Pharm Sci* **97**, 764–774

43. Mahler, H.-C. et al. Adsorption Behavior of a Surfactant and a Monoclonal Antibody to Sterilizing-Grade Filters. *J Pharm Sci* **99**, 2620–2627 (2010).

44. Bee, J. S., Randolph, T. W., Carpenter, J. F., Bishop, S. M. & Dimitrova, M. N. Effects of surfaces and leachables on the stability of biopharmaceuticals. *J Pharm Sci* **100**, 4158–4170 (2011).

45. Höger, K., Mathes, J. & Frieß, W. IgG1 adsorption to siliconized glass vials-influence of pH, ionic strength, and nonionic surfactants. *J Pharm Sci* **104**, 34–43 (2015).

46. Biddlecombe, J. G. et al. Determining antibody stability: creation of solid-liquid interfacial effects within a high shear environment. *Biotechnol Prog* **23**, 1218–1222 (2007).

47. Di Fede, G. et al. Monoclonal antibodies and antibody fragments: state of the art and future perspectives in the treatment of non-haematological tumors. *Expert Opin. Biol. Ther.* **11**, 1433–1445 (2011).

48. Saluja, A. & Kalonia, D. S. Nature and consequences of protein-protein interactions in high protein concentration solutions. *International Journal of Pharmaceutics* **358**, 1–15 (2008).

49. Torisu, T., Maruno, T., Hamaji, Y., Ohkubo, T. & Uchiyama, S. Synergistic Effect of Cavitation and Agitation on Protein Aggregation. *J. Pharm. Sci.* **106**, 521–529 (2017).

50. Rathore, N. et al. Characterization of protein rheology and delivery forces for combination products. *J. Pharm. Sci.* **101**, 4472–4480 (2012).

51. Ludwig, D. B., Carpenter, J. F., Hamel, J.-B. & Randolph, T. W. Protein adsorption and excipient effects on kinetic stability of silicone oil emulsions. *J Pharm Sci* **9999**, n/a (2009).

52. Basu, P., Blake-Haskins, A. W., O'Berry, K. B., Randolph, T. W. & Carpenter, J. F. Albinterferon α2b adsorption to silicone oil-water interfaces: Effects on protein conformation, aggregation, and subvisible particle formation. *J. Pharm. Sci.* **103**, 427–436 (2014).

53. Gerhardt, A. et al. Protein aggregation and particle formation in prefilled glass syringes. *J Pharm Sci* **103**, 1601–1612 (2014).

54. Thirumangalathu, R. et al. Silicone oil- and agitation-induced aggregation of a monoclonal antibody in aqueous solution. *J Pharm Sci* **98**, 3167–3181 (2009).

55. Kaur, K. Nanoemulsions as delivery vehicles for food and pharmaceuticals. in *Emulsions* 611–649 (Elsevier, 2016). doi:10.1016/B978-0-12-804306-6.00018-0

56. Gerhardt, A., Nguyen, B. H., Lewus, R., Carpenter, J. F. & Randolph, T. W. Effect of the siliconization method on particle generation in a monoclonal antibody formulation in pre-filled syringes. *J. Pharm. Sci.* **104**, 1601–1609 (2015).

57. Yano, Y. F. Kinetics of protein unfolding at interfaces. *Journal of Physics Condensed Matter* **24**, (2012).

58. Rabe, M., Verdes, D. & Seeger, S. Understanding protein adsorption phenomena at solid surfaces. *Adv Colloid Interface Sci* **162**, 87–106 (2011).
59. Pinholt, C., Hartvig, R. A., Medlicott, N. J. & Jorgensen, L. The importance of interfaces in protein drug delivery - why is protein adsorption of interest in pharmaceutical formulations? *Expert Opin Drug Deliv* **8**, 949–964 (2011).

60. Demanèche, S., Chapel, J. P., Monrozier, L. J. & Quiquampoix, H. Dissimilar pH-dependent adsorption features of bovine serum albumin and α-chymotrypsin on mica probed by AFM. *Colloids Surfaces B Biointerfaces* **70**, 226–231 (2009).

61. Rabe, M., Verdes, D., Zimmermann, J. & Seeger, S. Surface organization and cooperativity during nonspecific protein adsorption events. *J. Phys. Chem. B* **112**, 13971–13980 (2008).

62. Vroman, L. & Adams, A. L. Findings with the recording ellipsometer suggesting rapid exchange of specific plasma proteins at liquid/solid interfaces. *Surf. Sci.* **16**, 438–446 (1969).

63. Arai T, N. W. The behaviour of some model proteins at the solid-liquidinterface. *colloids surf* **51**, 1–15 (1990).

64. Jeon, S. I., Lee, J. H., Andrade, J. D. & De Gennes, P. G. Protein-surface interactions in the presence of polyethylene oxide. I. Simplified theory. *J. Colloid Interface Sci.* **142**, 149–158 (1991).

65. Jeon, S. I. & Andrade, J. D. Protein-surface interactions in the presence of polyethylene oxide. II. Effect of protein size. *J. Colloid Interface Sci.* **142**, 159–166 (1991).

66. Holmberg, M. & Hou, X. Competitive protein adsorption--multilayer adsorption and surface induced protein aggregation. *Langmuir* **25**, 2081–2089 (2009).

67. Kastantin, M., Langdon, B. B. & Schwartz, D. K. A Bottom-Up Approach to Understanding Protein Layer Formation at Solid-Liquid Interfaces. *Adv Colloid Interface Sci* **0**, 240–252 (2014).

68. Langmuir, I. Vapor pressures, evaporation, condensation and adsorption. *J. Am. Chem. Soc.* **54**, 2798–2832 (1932).

69. Cadilhe, A., Araújo, N. A. M. & Privman, V. Random sequential adsorption: From continuum to lattice and pre-patterned substrates. *J. Phys. Condens. Matter* **19**, (2007).

70. Norde, W. My voyage of discovery to proteins in flatland ...and beyond. *Colloids Surf B Biointerfaces* **61**, 1–9 (2008).

71. Kurrat, R., Ramsden, J. J. & Prenosil, J. E. Kinetic model for serum albumin adsorption: experimental verification. *J. Chem. Soc., Faraday Trans.* **90**, 587 (1994).

72. Tie, Y., Calonder, C. & Van Tassel, P. R. Protein adsorption: Kinetics and history dependence. *J Colloid Interface Sci* **268**, 1–11 (2003).

73. Chatelier, R. C. & Minton, A. P. Adsorption of globular proteins on locally planar surfaces: Models for the effect of excluded surface area and aggregation of adsorbed protein on adsorption equilibria. *Biophys. J.* **71**, 2367–2374 (1996).

74. Minton, A. Effects of excluded surface area and adsorbate clustering on surface adsorption of proteins. II. Kinetic models. *Biophys. J.* **80**, 1641–1648 (2001).

75. Sluzky, V., Tamada, J. A., Klibanov, A. M. & Langer, R. Kinetics of insulin aggregation in aqueous solutions upon agitation in the presence of hydrophobic surfaces. *Proc Natl Acad Sci U S A* **88**, 9377–9381 (1991).

76. Ballet, T. et al. DnaK prevents human insulin amyloid fiber formation on hydrophobic surfaces. *Biochemistry* **51**, 2172–2180 (2012).
77. Roberts, C. J. Therapeutic protein aggregation: Mechanisms, design, and control. *Trends in Biotechnology* **32**, 372–380 (2014).

78. Norde, W., MacRitchie, F., Nowicka, G. & Lyklema, J. Protein adsorption at solid-liquid interfaces: Reversibility and conformation aspects. *J. Colloid Interface Sci.* **112**, 447–456 (1986).

79. Norde, W. & Giacomelli, C. E. BSA structural changes during homomolecular exchange between the adsorbed and the dissolved states. in *Journal of Biotechnology* **79**, 259–268 (2000).

80. Vermonden, T., Giacomelli, C. E. & Norde, W. Reversibility of Structural Rearrangements in Bovine Serum Albumin during Homomolecular Exchange from Agl Particles. *Langmuir* **17**, 3734–3740 (2001).

81. Bee, J. S., Davis, M., Freund, E., Carpenter, J. F. & Randolph, T. W. Aggregation of a monoclonal antibody induced by adsorption to stainless steel. *Biotechnol. Bioeng.* **105**, 121–129 (2010).

82. Gurdak, E., Dupont-Gillain, C. C., Booth, J., Roberts, C. J. & Rouxhet, P. G. Resolution of the vertical and horizontal heterogeneity of adsorbed collagen layers by combination of QCM-D and AFM. *Langmuir* **21**, 10684–10692 (2005).

83. Herrera Estrada, L. P. & Champion, J. A. Protein nanoparticles for therapeutic protein delivery. *Biomater. Sci.* **3**, 787–799 (2015).

84. Reuter, B. & Petersen, C. Syringe siliconisation: Trends, methods, analysis procedures. *ONdrugDelivery* **16–19** (2013).

85. Krayukhina, E., Tsumoto, K., Uchiyama, S. & Fukui, K. Effects of syringe material and silicone oil lubrication on the stability of pharmaceutical proteins. *J. Pharm. Sci.* **104**, 527–535 (2015).

86. Hickel, A., Radke, C. J. & Blanch, H. W. Role of organic solvents on Pa-hydroxynitrile lyase interfacial activity and stability. *Biotechnol. Bioeng.* **74**, 18–28 (2001).

87. Zambaux, M. F., Bonneaux, F., Gref, R., Dellacherie, E. & Vigneron, C. Preparation and characterization of protein C-loaded PLA nanoparticles. *J. Control. Release* **60**, 179–188 (1999).

88. Gurkov, T. D., Russev, S. C., Danov, K. D., Ivanov, I. B. & Campbell, B. Monolayers of globular proteins on the air/water interface: Applicability of the volmer equation of state. *Langmuir* **19**, 7362–7369 (2003).

89. Beverung, C. J., Radke, C. J. & Blanch, H. W. Protein adsorption at the oil/water interface: Characterization of adsorption kinetics by dynamic interfacial tension measurements. *Biophys. Chem.* **81**, 59–80 (1999).

90. Koepf, E., Eisele, S., Schroeder, R., Brezesinski, G. & Friess, W. Notorious but not understood: How liquid-air interfacial stress triggers protein aggregation. *Int. J. Pharm.* **537**, 202–212 (2018).

91. Wrobel, S. Bubbles, Babies and Biology: The Story of Surfactant. *FASEB J.* **18**, 1624e–1624e (2004).

92. Pérez-Gil, J. Structure of pulmonary surfactant membranes and films: The role of proteins and lipid-protein interactions. *Biochim. Biophys. Acta - Biomembr.* **1778**, 1676–1695 (2008).
93. Lad, M. D., Birembaut, F., Matthew, J. M., Frazier, R. A. & Green, R. J. The adsorbed conformation of globular proteins at the air/water interface. *Phys. Chem. Chem. Phys.* **8**, 2179 (2006).

94. Jayaraman, M., Buck, P. M., Alphonse Ignatius, A., King, K. R. & Wang, W. Agitation-induced aggregation and subvisible particulate formation in model proteins. *Eur. J. Pharm. Biopharm.* **87**, 299–309 (2014).

95. Liu, L., Qi, W., Schwartz, D. K., Randolph, T. W. & Carpenter, J. F. The effects of excipients on protein aggregation during agitation: An interfacial shear rheology study. *J. Pharm. Sci.* **102**, 2460–2470 (2013).

96. Ghazvini, S., Kalonia, C., Volkin, D. B. & Dhar, P. Evaluating the Role of the Air-Solution Interface on the Mechanism of Subvisible Particle Formation Caused by Mechanical Agitation for an IgG1 mAb. *J. Pharm. Sci.* **105**, 1643–1656 (2016).

97. Bee, J. S., Schwartz, D. K., Trabelsi, S., Randolph, T. W. & Stevenson, J. L. Production of particles of therapeutic proteins at the air-water interface during compression/dilation cycles. *Soft Matter* **8**, 10329–10335 (2012).

98. Rudiuk, S., Huille, S., Tribet, C. & Cohen-Tannoudji, L. Importance of the dynamics of adsorption and of a transient interfacial stress on the formation of aggregates of IgG antibodies. *Soft Matter* **8**, 2651–2661 (2012).

99. Frachon, T., Bruckert, F., Le Masne, Q., Monnin, E. & Weidenhaupt, M. Insulin Aggregation at a Dynamic Solid-Liquid-Air Triple Interface. *Langmuir* **32**, 13009–13019 (2016).

100. Migliorini, E., Weidenhaupt, M. & Catherine, P. A practical guide to characterize biomolecule adsorption on solid surfaces. *Biointerphases* **13** (6), in press (2018).

101. Khan, T. A., Mahler, H.-C. & Kishore, R. S. K. Key interactions of surfactants in therapeutic protein formulations: A review. *Eur. J. Pharm. Biopharm.* **97**, 60–67 (2015).

102. Kamerzell, T. J., Esfandiary, R., Joshi, S. B., Middaugh, C. R. & Volkin, D. B. Protein-excipient interactions: Mechanisms and biophysical characterization applied to protein formulation development. *Advanced Drug Delivery Reviews* **63**, 1118–1159 (2011).

103. Lee, H. J., McAuley, A., Schilke, K. F. & McGuire, J. Molecular origins of surfactant-mediated stabilization of protein drugs. *Adv. Drug Deliv. Rev.* **63**, 1160–1171 (2011).

104. Das, T. K. Protein Particulate Detection Issues in Biotherapeutics Development—Current Status. *AAPS PharmSciTech* **13**, 732–746 (2012).

105. Elder, D. P., Kuentz, M. & Holm, R. Pharmaceutical excipients - Quality, regulatory and biopharmaceutical considerations. *Eur. J. Pharm. Sci.* **87**, 88–99 (2016).

106. Wang, W. Advanced protein formulations. *Protein Science* **24**, 1031–1039 (2015).

107. Frokjaer, S. & Otzen, D. E. Protein drug stability: a formulation challenge. *Nat Rev Drug Discov* **4**, 298–306 (2005).

108. Veronese, F. Peptide and protein PEGylation: a review of problems and solutions. *Biomaterials* **22**, 405–417 (2001).

109. Kurtzhals, P. Pharmacology of Insulin Detemir. *Endocrinol. Metab. Clin. North Am.* **36**, 14–20 (2007).

110. Nault, L. et al. Human insulin adsorption kinetics, conformational changes and amyloidal aggregate formation on hydrophobic surfaces. *Acta Biomater* **9**, 5070–5079 (2013).
111. Nault, L., Vendrely, C., Brechet, Y., Bruckert, F. & Weidenhaupt, M. Peptides that form beta-sheets on hydrophobic surfaces accelerate surface-induced insulin amyloidal aggregation. *FEBS Lett* **587**, 1281–1286 (2013).

112. Chouchane, K. et al. Dual Effect of (LK)nL Peptides on the Onset of Insulin Amyloid Fiber Formation at Hydrophobic Surfaces. *J Phys Chem B* **119**, 10543–10553 (2015).

113. Chouchane, K., Pignot-Paintrand, I., Bruckert, F. & Weidenhaupt, M. Visible light-induced insulin aggregation on surfaces via photoexcitation of bound thioflavin T. *J. Photochem. Photobiol. B Biol.* **181**, 89–97 (2018).

114. Timofeev, V. I. et al. X-ray investigation of gene-engineered human insulin crystallized from a solution containing polysialic acid. *Acta Crystallogr. Sect. F Struct. Biol. Cryst. Commun.* **66**, 259–263 (2010).

115. Murray-Rust, J., McLeod, A. N., Blundell, T. L. & Wood, S. P. Structure and evolution of insulins: Implications for receptor binding. *BioEssays* **14**, 325–331 (2018).

116. Huus, K., Havelund, S., Olsen, H. B., van de Weert, M. & Frokjaer, S. Thermal dissociation and unfolding of insulin. *Biochemistry* **44**, 11171–11177 (2005).

117. Jeffrey, P. D., Milthorpe, B. K. & Nichol, L. W. Polymerization pattern of insulin at pH 7.0. *Biochemistry* **15**, 4660–4665 (1976).

118. Milthorpe, B. K., Nichol, L. W. & Jeffrey, P. D. The polymerization pattern of zinc(II)-insulin at pH 7.0. *BBA - Protein Struct.* **495**, 195–202 (1977).

119. Waugh, D. F. A Fibrous Modification of Insulin. I. The Heat Precipitate of Insulin. *J. Am. Chem. Soc.* **68**, 247–250 (1946).

120. Irsigler, K. & Kritz, H. Long-term continuous intravenous insulin therapy with a portable insulin dosage-regulating apparatus. *Diabetes* **28**, 196–203 (1979).

121. Chantelau, E. A. & Berger, M. Pollution of insulin with silicone oil, a hazard of disposable plastic syringes. *Lancet* **1**, 1459 (1985).

122. Bernstein, R. K. Clouding and deactivation of clear (regular) human insulin: association with silicone oil from disposable syringes? *Diabetes Care* **10**, 786–787 (1987).

123. Thurow, H. & Geisen, K. Stabilisation of dissolved proteins against denaturation at hydrophobic interfaces. *Diabetologia* **27**, 212–218 (1984).

124. Sluzky, V., Klibanov, A. M. & Langer, R. Mechanism of insulin aggregation and stabilization in agitated aqueous solutions. *Biotechnol Bioeng* **40**, 895–903 (1992).

125. Ballet, T. Stability of therapeutic proteins in contact with materials: Surface-induced aggregation and detection of conformational changes by molecular chaperones. (2010).

126. Nakanishi, K., Sakiyama, T. & Imamura, K. On the adsorption of proteins on solid surfaces, a common but very complicated phenomenon. *J. Biosci. Bioeng.* **91**, 233–244 (2001).

127. Dzwolak, W., Smirnovas, V., Jansen, R. & Winter, R. Insulin forms amyloid in a strain-dependent manner: an FT-IR spectroscopic study. *Protein Sci* **13**, 1927–1932 (2004).

128. Jorgensen, L. et al. Adsorption of insulin with varying self-association profiles to a solid Teflon surface - Influence on protein structure, fibrillation tendency and thermal stability. *Eur. J. Pharm. Sci.* **42**, 509–516 (2011).

129. Mauri, S., Volk, M., Byard, S., Berchtold, H. & Arnolds, H. Stabilization of Insulin by Adsorption on a Hydrophobic Silane Self-Assembled Monolayer. *Langmuir* **31**, 8892–8900 (2015).
130. Sharp, J. S., Forrest, J. A. & Jones, R. A. Surface denaturation and amyloid fibril formation of insulin at model lipid-water interfaces. *Biochemistry* **41**, 15810–15819 (2002).

131. Smith, M. I., Sharp, J. S. & Roberts, C. J. Nucleation and growth of insulin fibrils in bulk solution and at hydrophobic polystyrene surfaces. *Biophys J* **93**, 2143–2151 (2007).

132. Jorgensen, L. et al. Adsorption of insulin with varying self-association profiles to a solid Teflon surface--influence on protein structure, fibrillation tendency and thermal stability. *Eur J Pharm Sci* **42**, 509–516 (2011).

133. Pandey, L. M. et al. Surface chemistry at the nanometer scale influences insulin aggregation. *Colloids Surf B Biointerfaces* **100**, 69–76 (2012).

134. Hosseinzadeh, G., Maghari, A., Farnia, S. M. F. & Moosavi-Movahedi, A. A. Interaction mechanism of insulin with ZnO nanoparticles by replica exchange molecular dynamics simulation. *J. Biomol. Struct. Dyn.* **1102**, 1–13 (2017).

135. Grigolato, F., Colombo, C., Ferrari, R., Rezabkova, L. & Arosio, P. Mechanistic Origin of the Combined Effect of Surfaces and Mechanical Agitation on Amyloid Formation. *ACS Nano* **11**, 11358–11367 (2017).

136. Ivanova, M. I., Sievers, S. A., Sawaya, M. R., Wall, J. S. & Eisenberg, D. Molecular basis for insulin fibril assembly. *Proc Natl Acad Sci U S A* **106**, 18990–18995 (2009).

137. DeGrado Lear JD, W. Induction of peptide conformation at apolar/water interfaces. 1. A study with model peptides of defined hydrophobic periodicity. *J. Am. Chem. Soc.* **107**, 7684–7689 (1985).

138. Chouchane, K. *Controlling surface mediated insulin aggregation by peptides and light*. (2017).

139. Cohen, S. I. A., Vendruscolo, M., Dobson, C. M. & Knowles, T. P. J. From macroscopic measurements to microscopic mechanisms of protein aggregation. *Journal of Molecular Biology* **421**, 160–171 (2012).

140. Gleason, C., Yee, C., Masatani, P., Middaugh, C. R. & Vance, A. Probing Shear Thinning Behaviors of IgG Molecules at the Air-Water Interface via Rheological Methods. *Langmuir* **32**, 496–504 (2016).

141. Wiesbauer, J., Prassl, R. & Nidetzky, B. Renewal of the air-water interface as a critical system parameter of protein stability: aggregation of the human growth hormone and its prevention by surface-active compounds. *Langmuir* **29**, 15240–15250 (2013).

142. Vieillard, V. et al. Pneumatic conveying systems and physical stability of monoclonal antibodies : example of Cetuximab. in *ECCO congress* poster (2013).

143. Sergii Rudiuk Sylvain Huille, Christophe Tribet, L. C.-T. Importance of the dynamics of adsorption and of a transient interfacial stress on the formation of aggregates of IgG antibodies. *Softmatter* **8**, 2651–2661 (2012).

144. Wang, S. et al. Stabilizing two IgG1 monoclonal antibodies by surfactants: Balance between aggregation prevention and structure perturbation. *Eur. J. Pharm. Biopharm.* **114**, 263–277 (2017).

145. Khan, T. A., Mahler, H.-C. & Kishore, R. S. K. Key interactions of surfactants in therapeutic protein formulations: A review. *Eur. J. Pharm. Biopharm.* **97**, Part A, 60–67 (2015).

146. Garidel, P., Hoffmann, C. & Blume, A. A thermodynamic analysis of the binding interaction between polysorbate 20 and 80 with human serum albumins and
immunoglobulins: A contribution to understand colloidal protein stabilisation. *Biophys. Chem.* **143**, 70–78 (2009).

147. Kapp, S. J., Larsson, I., Van De Weert, M., Cárdenas, M. & Jorgensen, L. Competitive Adsorption of Monoclonal Antibodies and Nonionic Surfactants at Solid Hydrophobic Surfaces. *J Pharm Sci* **104**, 593–601

148. Serno, T., Härtl, E., Besheer, A., Miller, R. & Winter, G. The Role of Polysorbate 80 and HPβCD at the Air-Water Interface of IgG Solutions. *Pharm Res* **30**, 117–130 (2013).

149. Gerhardt, A. et al. Surfactant Effects on Particle Generation in Antibody Formulations in Pre-filled Syringes. *J Pharm Sci* **104**, 4056–4064 (2015).

150. Mackie, A. R., Gunning, A. P., Wilde, P. J. & Morris, V. J. Orogenic Displacement of Protein from the Air/Water Interface by Competitive Adsorption. *J Colloid Interface Sci* **210**, 157–166 (1999).

151. Shortall, A. ADVANCED DELIVERY DEVICES - The Reconstitution Revolution - *Drug Dev. Deliv.* (2013).

152. Bee, J. S. et al. Monoclonal antibody interactions with micro- and nanoparticles: adsorption, aggregation, and accelerated stress studies. *J Pharm Sci* **98**, 3218–3238 (2009).

153. Buijs, J., Lichtenbelt, J. W. T., Norde, W. & Lyklema, J. Adsorption of monoclonal IgGs and their F(ab')2 fragments onto polymeric surfaces. *Colloids Surfaces B Biointerfaces* **5**, 11–23 (1995).

154. Wiseman, M. E. & Frank, C. W. Antibody Adsorption and Orientation on Hydrophobic Surfaces. *Langmuir* **28**, 1765–1774 (2012).

155. Bee, J. S. et al. Monoclonal antibody interactions with micro- and nanoparticles: adsorption, aggregation, and accelerated stress studies. *J Pharm Sci* **98**, 3218–3238 (2009).

156. Vermeer, A. W. P., Giacomelli, C. E. & Norde, W. Adsorption of IgG onto hydrophobic teflon. Differences between the Fab and Fc domains. *Biochim. Biophys. Acta - Gen. Subj.* **1526**, 61–69 (2001).

157. Joshi, O. & McGuire, J. Adsorption behavior of lysozyme and Tween 80 at hydrophilic and hydrophobic silica-water interfaces. *Appl Biochem Biotechnol* **152**, 235–248 (2009).

158. Joshi, O., McGuire, J. & Wang, D. Q. Adsorption and function of recombinant factor VIII at solid&#x2192;water interfaces in the presence of Tween-80. *J Pharm Sci* **97**, 4741–4755

159. Couston, R. G., Skoda, M. W., Uddin, S. & van der Walle, C. F. Adsorption behavior of a human monoclonal antibody at hydrophilic and hydrophobic surfaces. *MAbs* **5**, 126–139 (2013).

160. Schuck, P. & Zhao, H. The Role of Mass Transport Limitation and Surface Heterogeneity in the Biophysical Characterization of Macromolecular Binding Processes by SPR Biosensing. *Methods Mol. Biol.* **627**, 15–54 (2010).

161. Smoluchowski, M. v. Versuch einer mathematischen Theorie der Koagulationskinetik kolloiderLösnnngen. *Kolloid-Zeitschrift* **21**, 98–104 (1917).

162. Hall, D. R., Cann, J. R. & Winzor, D. J. Demonstration of an upper limit to the range of association rate constants amenable to study by biosensor technology based on surface plasmon resonance. *Anal. Biochem.* **235**, 175–184 (1996).
163. Kim, H. L., Mcauley, A. & Mcguire, J. Protein Effects on Surfactant Adsorption Suggest the Dominant Mode of Surfactant-Mediated Stabilization of Protein. *J. Pharm. Sci.* **103**, 1337–1345 (2014).

164. Ausserré, D. & Valignat, M.-P. Surface enhanced ellipsometric contrast (SEEC) basic theory and lambda/4 multilayered solutions. *Opt. Express* **15**, 8329–8339 (2007).

165. Souplet, V., Desmet, R. & Melnyk, O. Imaging of protein layers with an optical microscope for the characterization of peptide microarrays. *J Pept Sci* **13**, 451–457 (2007).

166. Ducret, A., Valignat, M. P., Mouhamar, F., Mignot, T. & Theodoly, O. Wet-surface-enhanced ellipsometric contrast microscopy identifies slime as a major adhesion factor during bacterial surface motility. *Proc Natl Acad Sci U S A* **109**, 10036–10041 (2012).

167. Xi, W.-H. & Wei, G.-H. Amyloid- β peptide aggregation and the influence of carbon nanoparticles. *Chinese Phys. B* **25**, 18704 (2016).

168. Roach, P., Farrar, D. & Perry, C. C. Interpretation of protein adsorption: surface-induced conformational changes. *J Am Chem Soc* **127**, 8168–8173 (2005).

169. Jichao Kang, Jason Penera, X. L. Rapid Formulation Development for Monoclonal Antibodies. *Bioprocess Int.* **14**, (2016).

170. Sluzky, V., Langer, R. & Klibanov, A. M. Insulin stability and aggregation in agitated aqueous solutions. *Department of Chemical Engineering PhD*, (Massachusetts Institute of Technology, 1992).
# 8 Annexe 1

List of students, other than PhD students, I supervised since 2010:

## M2R Master Theses

2011

J. Bhagyashree. Master Nanobiology, UJF Grenoble; Protein aggregation on hydrophobic surfaces by SPRi

S. Menad. Master Nanobiology, UJF Grenoble; Capillary force assembly of DNA: deposition, stretching and AFM structure.

2012

G. Kaur Behrh, Master Nanobiology, UJF Grenoble; Adsorption kinetics of therapeutic proteins on material surfaces.

2013

K. Chouchane, Master Nanobiology, UJF Grenoble; Effect of peptides on insulin aggregation kinetics in contact with hydrophobic material surfaces.

2016

G Lefebvre, Master Sciences Pharmaceutiques "Nanomédecines", Univ Angers; Stability study of a therapeutic monoclonal antibody at the liquid-solid interface.

## M1 internships

2010

C. Monnot (stage 2A Phelma SMPB); Etude de l'étalement de mutants de Disctyostelium discoideum sur des surfaces

2011

G. Bouchage (stage 2A Phelma SMPB); iGEM 2011: MercuroColi : a new way to quantify heavy metals

2012

A. Millet (stage M1 Biologie Moléculaire et Cellulaire UJF); iGEM 2012: sEnsiColi: A tunable and reliable ultra-sensitive detector

2013

A. Rapeaux (stage 2A Phelma SMPB); iGEM 2013: Talk E'coli
2014

A. Dolle (Stage ingénieur, Telecom Physique, Strasbourg); Agrégation de l'insuline en fibres amyloïdes sur des nanobilles fonctionnalisées

2017

S Cambronero (stage 2A Phelma Biomedical Engineering); Characterization of a drug in EVEON's fluidic technology

2018

X Gallard (stage 2A Phelma Biomedical Engineering); iGEM 2018 Phag'zer

BTS internships and international exchange students

2010

L. Lacroix, BTS AnaBiotech (analyses agricoles biologiques et biotechnologique), Lycée des Mandailles

V. Deroux, BTS AnaBiotech (analyses agricoles biologiques et biotechnologique), Lycée des Mandailles

2011

M. Norton, first year PhD student Summer Programm Minatec, UPenn, USA

P. Guo (Canada), third year bachelor student, Programme ORA Rhône-Alpes, Toronto University, Canada

2012

S. Fairley (Giant international internship program, Alabama State Univ, Montgomery, USA)

2013

K. Moreaux (Giant international internship program, Louisiana State Univ, Baton Rouge, USA)

2014

S. Humble (Giant international internship program, Louisiana State Univ, Baton Rouge, USA)

2017

B Weiner (Giant international internship program, UPenn, Philadelphia, USA)

S Barrow (Giant international internship program, Alabama State Univ, Montgomery, USA)

T Niepa (Giant international internship program, UPenn, Philadelphia, USA)

2018

K Rose (Giant international internship program, UPenn, Philadelphia, USA)
## 9 Annexe 2

List of oral and poster communications since 2010:

### Oral Communications (speaker in italics)

2010, IBC's 10th Annual, Formulation Strategies for Protein Therapeutics, Providence (RI) US.
F Bruckert, *T. Ballet*, P. Mangiagalli, C. Bureau, **M. Weidenhaupt**, Role of material surfaces in inducing protein aggregation.

2011, XXI International Symposium on Bioelectrochemistry and Bioenergetics, Krakow (Po),
*Nguyen U.H.P.*, Stambouli, V., **Weidenhaupt, M.**, *Delabouglise, D*, *Extracellular Redox Compounds produced by E. coli*

2011, Journées d'Electrochimie 2011, Grenoble (F), *Nguyen U.H.P.*, Stambouli, V., **Weidenhaupt, M.**, *Delabouglise, D*, *Extracellular Redox Compounds produced by E. coli*

2012, 13ème colloque du Groupe Français de Bioélectrochimie, Lacaneau (F), *Nguyen U.*, **Weidenhaupt, M.**, Geiselmann, H., Stambouli, V., D. Delabouglise, Electroactive deposit from Escherichia coli.

2013, Frontiers in BioMagnetic Particles, Telluride (Co) US, Zanini L.F., Blaire, G., Le Roy, D., Reyne, G., Cugat, O., **Weidenhaupt, M.**, Bruckert, F., Dumas-Bouchiat, F., *Dempsey, N.M.*
Development of hard magnetic micropatterns for biomedical applications.

2015, Biophysics of proteins at Surfaces, Madrid (E), *Chouchane K, Amari, M*, Vendrely C, *Moreaux K*, Bruckert F, **Weidenhaupt M**, Peptides forming beta-sheets on hydrophobic surfaces cooperatively promote insulin amyloidal aggregation.

2017, IMAPS, Lyon (F), *T Frachon*, F Bruckert, R Vomscheid, H Marko, E Monnin, Q LeMasne, **M Weidenhaupt**, Therapeutic protein aggregation at the triple interface air/liquid/solid: relevance to drug production, storage and delivery processes

### Poster communications (student underlined)

2010 Workshop on Protein Aggregation and Immunogenicity, Breckenridge (Co), US.
Mangiagalli F, Felsovalyi, *T. Ballet*, F. Bruckert, **M. Weidenhaupt**, S. Kumar, S. Banta, C. Bureau.
Development of pharmaceutically relevant model systems to study the role of solid surfaces
in inducing protein aggregation and conformational changes.

2010 PEGS (Protein Aggregation in Biopharmaceutical Products), Boston (Ma) US. Mangiagalli
T. Ballet, F. Felsovalyi, F. Bruckert, **M. Weidenhaupt**, S. Kumar, S. Banta, C. Bureau, P.
Evaluating the role of solid surfaces in Inducing Aggregation and Conformational Changes in
Model Proteins.

2010 4th annual seminar of the Cluster de Recherche Macodev, Annecy (F). Nault L., Ballet, T., **Weidenhaupt, M.**, Bruckert, F., Brechet, Y. Mechanism of amyloïd fiber formation on material surfaces.

2010 The 61st annual meeting of the International Society of Electrochemistry, Nice (F),
**Nguyen U.H.P.**, Stambouli, V., Delabouglise, D., **Weidenhaupt, M.**, Douki, T. Electrochemistry
of Bacterial Extracellular Redox Compounds.

2010, The Physical Cell, London(UK). Socol M., Lefrou, C., Bruckert, F., Delabouglise, D., **Weidenhaupt, M.** Electrochemically-triggered, synchronized adhesion and spreading of Dictyostelium discoideum.

2010, WCARP IV, Arcachon (F). Socol M., Lefrou, C., Bruckert, F., Delabouglise, D., **Weidenhaupt, M.** Electrochemically-triggered, synchronized adhesion and spreading of Dictyostelium discoideum.

2011, FEBS, Torino (I). Nault L., Ballet, T., Bruckert, F., Brechet, Y., **Weidenhaupt, M.** Protein stability upon contact with material surfaces.

2011, 5th annual seminar of the Cluster de Recherche Macodev, Annecy (F), L. Nault, T. Ballet, Y. Brechet, F. Bruckert, **M. Weidenhaupt** Mécanisme d'agrégation des protéines en contact avec les surfaces hydrophobes.

2012, 4th Workshop on Nano and Micro-Environments for Cell Biology, Grenoble (F) Blaire G., Masse, A., Zanini, L., Delshadi, S., Honegger, T., Bruckert, F., Peyrade, D., **Weidenhaupt, M.**, Dempsey, N. M., Dumas-Bouchiat, F., Reyne, G., Cugat, O. Bio-Mag-MEMS combining magnetophoresis and dielectrophoresis for cell sorting.

2012, JEMS, Parma (I), Blaire G., Masse, A., Zanini, L., Honegger, T., Bruckert, F., Peyrade, D., **Weidenhaupt, M.**, Dempsey, N. M., Dumas-Bouchiat, F., Reyne, G., Cugat, O. Magnetic
microfluidic microsystems (Bio-Mag-MEMS) combining magnetophoresis and dielectrophoresis for cell sorting.

2013, ERC Biomim, Grenoble (F), Nault L., Chouchane K., Vendrely, C., Bruckert, F., **Weidenhaupt, M.** Peptides that form β-sheets on hydrophobic surfaces accelerate surface-induced insulin amyloidal aggregation

2013, SEP Conference, Paris (F), Zanini L.F., Blaire, G., Le Roy, D., Piveta, J., Osman, O., Tomba, C., Villard, C., Frénéa-Robin, M., Haddour, N., Reyne, G., Cugat, O., **Weidenhaupt, M.**, Bruckert, F., Dumas-Bouchiat, F., Dempsey, N.M., Autonomous separation devices based on micro-magnets.

2015, Colloque ADEBiotech Stabilité et formulation des protéines et des peptides, Romainville (F), Chouchane K, Amari, M, Vendrely C, Moreaux K, Bruckert F, **Weidenhaupt M**, Peptides forming beta-sheets on hydrophobic surfaces cooperatively promote insulin amyloidal aggregation.

2015, Colloque ADEBiotech Stabilité et formulation des protéines et des peptides, Romainville (F), Frachon T, LeMasne Q, Bruckert F, **Weidenhaupt M**, Fast insulin amyloid aggregation at the air-liquid-solid triple interface.
10 Reprints of articles

The following articles are reprinted hereafter:

Ballet, T. et al. DnaK prevents human insulin amyloid fiber formation on hydrophobic surfaces.
Biochemistry 51, 2172–2180 (2012).

Nault, L. et al. Human insulin adsorption kinetics, conformational changes and amyloidal aggregate formation on hydrophobic surfaces. Acta Biomater 9, 5070–5079 (2013).

Nault, L., et al. Peptides that form beta-sheets on hydrophobic surfaces accelerate surface-
induced insulin amyloidal aggregation. FEBS Lett 587, 1281–1286 (2013).

Chouchane, K. et al. Dual Effect of (LK)nL Peptides on the Onset of Insulin Amyloid Fiber Formation at Hydrophobic Surfaces. J Phys Chem B 119, 10543–10553 (2015).

Frachon, T., et al. Insulin Aggregation at a Dynamic Solid-Liquid-Air Triple Interface. Langmuir
32, 13009–13019 (2016).
# DnaK Prevents Human Insulin Amyloid Fiber Formation on Hydrophobic Surfaces

Thomas Ballet,†,‡ Franz Brukert,† Paolo Mangiagalli,‡ Christophe Bureau,‡ Laurence Boulangé,‡,§
Laurent Nault,† Thomas Perret,† and Marianne Weidenhaupt*,†

**ABSTRACT:** We have developed a multiwell-based protein aggregation assay to study the kinetics of insulin adsorption and aggregation on hydrophobic surfaces and to investigate the molecular mechanisms involved. Protein-surface interaction progresses in two phases: (1) a lag phase during which proteins adsorb and prefibrillar aggregates form on the material surface and (2) a growth phase during which amyloid fibers form and then are progressively released into solution. We studied the effect of three bacterial chaperones, DnaK, DnaJ, and ClpB, on insulin aggregation kinetics. In the presence of ATP, the simultaneous presence of DnaK, DnaJ, and ClpB allows good protection of insulin against aggregation. In the absence of ATP, DnaK alone is able to prevent insulin aggregation. Furthermore, DnaK binds to insulin adsorbed on hydrophobic surfaces. This process is slowed in the presence of ATP and can be enhanced by the cochaperone DnaJ. The peptide LVEALYL, derived from the insulin B chain, is known to promote fast aggregation in a concentration- and pH-dependent manner in solution. We show that it also shortens the lag phase for insulin aggregation on hydrophobic surfaces. As this peptide is also a known DnaK substrate, our data indicate that the peptide and the chaperone might compete for a common site during the process of insulin aggregation on hydrophobic surfaces.

<figure><img src="image_57.png" /></figure>

Adsorption of protein on material surfaces is of widespread importance in fields like cellular biology, pharmacology, and medicine. It can be quantitatively measured either biochemically after desorption or in situ by sensitive biophysical methods, like QCM-D or surface plasmon resonance (SPR). Protein adsorption is accompanied by conformational changes upon contact with the material surface, leading sometimes to protein aggregation. It is not easy to monitor these conformational changes on the material in a fast, convenient, and preferentially multiplexed assay. Albeit greatly sensitive, techniques like attenuated total reflectance Fourier transform infrared spectroscopy (ATR-FTIR) are not readily available and do not provide unambiguous information about the nature of the changes in protein folding.

Within cells, chaperones constitute a set of proteins whose function is to check the folding state of other proteins, and to refold them,¹⁻³ using ATP hydrolysis energy to drive protein disaggregation and renaturation. In *Escherichia coli*, Bukau and co-workers identified DnaK, DnaJ, and ClpB as the minimum set of bacterial chaperones needed to help the cell recover from a heat shock.⁴ DnaK is an ATPase specific for unfolded hydrophobic amino acid stretches. DnaJ forms a complex with DnaK and stimulates DnaK ATPase activity.⁵ ClpB is a hexameric ATPase that interacts with aggregated proteins and disaggregates them, in cooperation with DnaK and DnaJ.⁴,⁶,⁷ Furthermore, in *E. coli*, ~250 proteins are disaggregated and

refolded when DnaK, DnaJ, and ClpB are present, which supports the broadness of their protein substrates.⁴ Chaperones detect general conformational unfolding characteristics, rather than specific targets. In addition, many chaperones work in complexes; hence, chaperone combination is likely to enhance the recognition process. We therefore hypothesized that this protein family could provide new sensors for protein conformational changes at material surfaces.

We chose human insulin (HI) as a model because its aggregation has been studied under different physicochemical conditions.⁸⁻¹⁰ It was shown that HI aggregation is formulation-dependent and that insulin undoubtedly changes its conformation upon binding to hydrophobic surfaces, leading to the formation and release of amyloid fiber aggregates.¹¹ From a concentration dependence study, Sluzky et al. deduced that the HI monomer was the molecular species leading to aggregation.⁹ Using recombinant chaperones, we observe that different sets of them are able to prevent HI aggregation by binding preferentially to surface-bound insulin. Rüdiger et al.¹² have shown that DnaK binds to the insulin B chain peptide SHLVEALYLVCGER, and Ivanova et al.¹³ demonstrated that the shorter LVEALYL peptide is the minimal sequence that can

<footnote>†Laboratoire des Matériaux et du Génie Physique, Grenoble Institute of Technology, 3 parvis Louis Néel, 38016 Grenoble Cedex 1, France</footnote>

<footnote>‡Becton-Dickinson Pharmaceutical Systems, 11, Rue Aristide Bergès, 38801 Le Pont de Claix, France</footnote>
accelerate the lag time of insulin fiber formation at acidic pH in a concentration-dependent manner. We show that this same peptide also accelerates the lag time of insulin aggregation on hydrophobic surfaces and that bacterial chaperones are able to counterbalance this effect.

## EXPERIMENTAL PROCEDURES

If not otherwise stated, all chemicals were purchased from Sigma-Aldrich. Experiments were conducted in TBS [25 mM Tris-HCl (pH 7.4) and 125 mM NaCl]. HI (recombinant, expressed in yeast) solutions were prepared at 0.5 mg/mL (86 µM). All solutions were filtered (0.22 µm) before being used. The LVEALYL peptide was purchased from Genecust (Luxembourg) and dissolved in 20 mM NaOH at a concentration of 4.3 mM.

**Bacterial Chaperone Preparation and Activity Assay.**
The bacterial strains used to produce DnaK (Hsp70), ClpB (Hsp100), and DnaJ (Hsp40) were kindly provided by B. Bukau and A. Mogk (Zentrum für Molekulare Biologie Heidelberg, Universität Heidelberg, Heidelberg, Germany). His-tagged ClpB and DnaJ were purified using nickel-nitrilotriacetic acid (Ni-NTA) metal affinity chromatography columns (Qiagen) according to the manufacturer's instructions. DnaK was purified as described by Cegielska and Georgopoulos¹⁴ and McCarty and Walker,¹⁵ with minor modifications by Buchberger et al.¹⁶ The refolding activity of the purified proteins was controlled using the malate dehydrogenase (MDH) renaturation assay.⁶,¹⁷ The three proteins and ATP were essential for the refolding activity (2 ± 0.3 nM min⁻¹ at 1 µM DnaK, 0.2 µM DnaJ, and 1 µM ClpB).

**Insulin Aggregation Assays.** HI aggregation assays were conducted as eight replicates in plastic 96-well microplates. Polystyrene (Greiner Bio-One, contact angle of 85 ± 4.7°) or PEO-coated (Corning, contact angle of 3.5 ± 5.8°) microplates were used. In fluorescence assays, black polystyrene microplates were used (Nunc Nunclon Δ Surface). The plates were covered by plastic sheets, incubated at 37 °C, and shaken at 1200 rpm (Heidolph Titramax, 1.5 mm vibration orbit). At each time point, the solution was pipetted out of the microwells. Part of the solution was filtered to remove aggregated HI (100 nm cutoff). The wells were washed twice with 300 µL of TBS. The adsorbed HI fraction was desorbed with 100 µL of 5% SDS for a 1 h agitation at 37 °C. Negligible protein material remained on the surface thereafter. The total amounts of HI in solution (nonfiltered), soluble HI, and HI adsorbed in the wells were determined using the bicinchoninic acid (BCA) assay.¹⁸⁻²¹ In addition, turbidity (λ = 600 nm) or thioflavin T (ThT, 20 or 50 µM) fluorescence was directly measured in the wells. Free and bound forms of ThT were measured at a λ<sub>ex</sub> of 342 nm and a λ<sub>em</sub> of 430 nm and at a λ<sub>ex</sub> of 450 nm and a λ<sub>em</sub> of 482 nm, respectively,²² with a 5 nm excitation and emission slit (Tecan Infinite M1000).

**Kinetic Analysis.** The aggregation kinetics proceed in three phases: a lag phase, where the signal was not statistically different from the baseline (mean ± standard deviation), a linear growth phase, and a plateau phase. Experimentally, the lag time was defined by the intercept between the linear growth phase and the baseline. The growth rate was defined as the slope of the linear phase and the plateau as the maximal value attained. The parameters were calculated on individual kinetics corresponding to different samples, and the given statistics represent the average and standard deviation for each parameter.

**Protection of Insulin Aggregation by Bacterial Chaperones.** HI aggregation was monitored by turbidity or ThT fluorescence and confirmed by protein quantification after filtration at the end of the experiment. The TBS buffer solution was supplemented with 2 mM MgCl₂ (TBS-M), because Mg²⁺ ions are needed for the chaperone ATPase activity. Magnesium itself does not affect HI aggregation. All chaperones were stable in this buffer. The efficiency of chaperone protection was expressed as the lag time before the onset of HI aggregation. As Zn²⁺ influences the equilibrium of HI hexamer formation,²³ we tested the effect of Zn²⁺ addition (up to 10 µM) on HI aggregation: neither HI aggregation kinetics nor DnaK protection was modified.

**Chaperone Binding Assays.** Polystyrene microplates [96 wells, Greiner enzyme-linked immunosorbent assay (ELISA)] were incubated at 37 °C and 1200 rpm with insulin for various amounts of time and blocked with 10 mg/mL bovine serum albumin (BSA) in TBS-M (blocking buffer) for 30 min at room temperature (RT) with shaking (1000 rpm). DnaK, DnaJ, or both (diluted in blocking buffer) were then added for 30 min (RT) with shaking (1000 rpm). The plate was blocked again with BSA for 30 min (RT) with shaking (1000 rpm).

For DnaK detection, a mouse anti-DnaK (E. coli) monoclonal antibody (8E2/2; diluted 1:2500 in blocking buffer) was used, and for DnaJ detection, a mouse anti-penta-His antibody (diluted 1:1000 in blocking buffer) was added to each well and the mixture incubated for 15 min (RT) with shaking (1000 rpm). Finally, a goat horseradish peroxidase-conjugated anti-mouse antibody (diluted 1:2500 in blocking buffer) was added to each well and the mixture incubated for 15 min (RT) with shaking (1000 rpm). Wells were then washed three times with 200 µL of TBS, and 200 µL of ECL substrate solution was added per well. Chemiluminescence signals were immediately recorded using a TriStar LB 941 microplate multimode reader.

To relate luminescence values to the amount of DnaK adsorbed, a calibration experiment was conducted in parallel. Increasing amounts of DnaK were incubated in 96-well polystyrene microplates (Greiner ELISA). The amounts of adsorbed DnaK were determined using the NanoOrange assay, according to the manufacturer's instructions. An ELISA was performed on the adsorbed DnaK as described previously. A calibration curve was then obtained that related luminescence values to the amount of adsorbed DnaK.

## RESULTS

**HI Aggregation in the Presence of Hydrophobic Surfaces.** To study the mechanisms of insulin aggregation on hydrophobic surfaces, we developed a HI aggregation assay using commercial hydrophilic or hydrophobic 96-well plates, to allow rapid screening of different experimental conditions. A HI solution was placed in the presence of hydrophilic or hydrophobic plastic surfaces and agitated at 37 °C. At the indicated times, the solution was recovered and filtered to separate the soluble and aggregated HI pools. The microwell surface was then washed, and adsorbed HI was desorbed with SDS. These three protein pools were then quantified using the BCA assay. In the presence of hydrophilic surfaces, HI remained soluble for several days and less than 1 µg of protein was adsorbed on the hydrophilic microwell surfaces (data not shown). In contrast, in the presence of hydrophobic surfaces, the concentration of soluble HI remained constant for ~4 h (lag phase), after which the amount decreased
sharply to trace amounts (Figure 1A). The amount of aggregated HI in solution increased in parallel at a similar rate.

<figure><img src="image_58.png" /><figcaption>**Figure 1.** Kinetics of human insulin adsorption and aggregation in hydrophobic microplates. (A) Amount of soluble (●), aggregated (▲), and adsorbed (■) HI after the indicated incubation time. (B) Amyloid fiber-bound ThT fluorescence in solution (▲) and on the surface (■) plotted as a function of incubation time. The lag time as the intercept between the growth rate (—) and the baseline is denoted with an arrow. Other curves are hand-drawn and provided as a guide for the eye.</figcaption></figure>

The amount of HI adsorbed on hydrophobic surfaces increased during the aggregation process, reaching a maximum amount of ~40 µg.

Amyloidal aggregates are characterized by the formation of intermolecular β-sheets, which can be probed by the binding of thioflavin T (ThT), resulting in a characteristic fluorescence signal. Both the aggregated and the adsorbed HI pools were stained with ThT, and the fluorescence intensity per microgram of protein was the same in both pools (Figure 1B). The ThT fluorescence is therefore a convenient method for monitoring the appearance of HI aggregates. In the following experiments, we defined the lag time as the intercept between the linear growth phase and the baseline of the ThT fluorescence kinetics. Depending on HI preparations, the lag time varied from 2 to 4 h. Within an experiment, using the same HI preparation in the same multiwell plate, the lag time was also variable from well to well, which explains the variation observed in the aggregation kinetics. HI aggregates exhibited elongated fiberlike rods with a diameter of 5–10 nm and a length of 50–100 nm when imaged using electron microscopy (data not shown). Their morphology and dimensions are similar to those of the fibers obtained after incubation of insulin at pH 2 and an elevated temperature.²⁴,²⁵

**Effect of Bacterial Chaperones on Insulin Aggregation.** We then studied whether bacterial chaperones could have an effect on the kinetics of HI aggregation. In the presence of 1 mM ATP, the simultaneous presence of the three chaperones delayed HI aggregation as shown by the 9-fold increase in the

aggregation lag time (Figure 2A). DnaK alone slightly increased the lag time, whereas DnaJ was without effect. The combined

<figure><img src="image_59.png" /><figcaption>**Figure 2.** Effect of bacterial chaperones on human insulin aggregation. A HI solution was agitated in a hydrophobic microplate, in the presence of DnaK (0.3 µM), DnaJ [0.06 µM (+) and 0.3 µM (++)], and ClpB [0.3 µM (+) and 0.6 µM (++)] as indicated (A) in the presence of 1 mM ATP or (B) in the absence of ATP. The aggregation kinetics were monitored by turbidity, and the lag times were determined as explained in Experimental Procedures.</figcaption></figure>

presence of DnaK and DnaJ significantly increased the lag time, to a level similar to that obtained in the presence of ClpB alone. Moreover, the combined presence of ClpB with DnaK, but not DnaJ, further delayed HI aggregation in the presence of 1 mM ATP. When the concentrations of DnaJ and ClpB were increased (++ vs +), the lag time further increased. These results show that HI undergoes conformational changes during the aggregation process that are recognized and could possibly be repaired by a minimal set of bacterial chaperones. Nevertheless, incubation of 150 µg of final amyloidal insulin aggregates with DnaK, DnaJ, and ClpB for 24 h does not allow recovery of more than 1 µg of soluble insulin (data not shown). This suggests that the three chaperones do not dissociate final aggregates but more likely block intermediate aggregation states.

Surprisingly, in the absence of ATP, no protein aggregation was observed whenever DnaK was present (Figure 2B). The effect was specific for DnaK because, in the absence of ATP, the presence of the other chaperones alone or in combination did not significantly change the HI aggregation kinetics. Addition of comparable and larger amounts of BSA, which is routinely used as a blocking agent to cover material surfaces and works by competitive adsorption, had no effect (data not shown). The peculiar role of DnaK is further investigated in the next section.

**Prevention of Insulin Aggregation by DnaK in the Absence of ATP.** In the presence of 1 mM ATPγS, a
nonhydrolyzable form of ATP, DnaK did not prevent HI aggregation (Figure 3A), which showed that no ATP hydrolysis

<figure><img src="image_60.png" /></figure>

**Figure 3.** Nucleotide requirements for DnaK protection of human insulin aggregation on hydrophobic surfaces. A HI solution was agitated in a hydrophobic microplate in the presence of the indicated chaperones and nucleotides. HI aggregation was monitored by optical density measurements at 600 nm. (A) The HI solution was supplemented with 1 µM DnaK and either no nucleotide (○) or 1 mM ATP, ADP, or ATPγS (●, ▲, and ■, respectively). (B) The HI solution was supplemented with 0.3 µM DnaK, and the aggregation rate is plotted as a function of ATP concentration. (C) The HI solution was not supplemented with chaperones (♦), supplemented with 0.3 µM DnaK in the absence of ATP (△) or with 1 mM ATP (×), or supplemented with 0.3 µM DnaK and 0.06 µM DnaJ with 1 mM ATP (□).

took place on the HI substrate. Furthermore, addition of 1 mM ADP also abolished the protective effect of DnaK on HI aggregation. The similar effect obtained by the presence of ADP, ATP, or a nonhydrolyzable nucleotide suggested that the molecular species, active at preventing the formation of HI amyloidal fibers, was the nucleotide-free DnaK<sub>empty</sub>–HI complex. This fits well with the biochemical properties of this protein because in the presence of ATP, DnaK indeed has a decreased affinity for its protein substrate<sup>26,27</sup> and hence could be less protective against aggregation. A series of HI aggregation experiments were performed in the presence of DnaK at different ATP concentrations ranging from 1 µM to 5 mM. The HI aggregation rate was used to measure the effect of added ATP on DnaK (Figure 3B). An apparent affinity of 2 µM was determined, which fits with the reported values of the affinity of DnaK for ATP.<sup>28</sup> Furthermore, in the presence of ATP, DnaJ

increased the protective effect of DnaK on HI aggregation (Figure 3C). It is indeed known that the DnaK–DnaJ complex has a higher affinity for the protein substrate than DnaK alone.<sup>29,30</sup>

Several lines of evidence indicate that DnaK prevented early phases of HI aggregation. First, in the continuous presence of DnaK, the amount of HI that adsorbed on hydrophobic surfaces remained small (Figure 4A) and ThT did not stain the adsorbed HI (Figure 4B, vs Figure 1B). Second, in two-stage experiments, when the protective effect of DnaK was released after 5 h by addition of ATP, HI aggregation took place after the same lag time observed in the absence of DnaK (Figure 4C). Similarly, when DnaK was added after incubation for 1 h, the lag time was reduced by 1 h after the addition of ATP. This showed that, as long as it was present in a nucleotide-free form, DnaK prevented the formation of amyloid aggregates on hydrophobic surfaces. To confirm the early effect of DnaK, a constant amount of DnaK was added at different times after the beginning of the incubation. After a 1 h pre-incubation, DnaK considerably slowed HI aggregation kinetics, but after 2 h, it had no significant effect, although aggregation had not yet started (Figure 4D).

### DnaK Binds to HI Adsorbed on Hydrophobic Surfaces.

We then determined the minimal amount of DnaK that should be added as a function of preincubation time to prevent HI aggregation for at least 18 h (Figure 5). This amount increased very rapidly without exceeding the amount of HI adsorbed on the surface, on a molar ratio basis.

Because insulin aggregates are present both on the plastic surface and in solution, we studied the binding of DnaK to the HI pool adsorbed on the surface and to HI aggregates in suspensions. BCA and sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) analysis were combined to quantify the amount of DnaK and HI adsorbed on the surface or to aggregates in suspension (Table 1). For these experiments, we used an experimental setup similar to that of Sluzky et al.<sup>8</sup> SurfaSil-treated borosilicate beads were incubated with HI under agitation overnight. After the beads had been washed, 20 ± 2 µg of HI remained adsorbed to the beads. The HI solution fully aggregated, and 150 µg of HI aggregates was recovered by centrifugation and washed.

In a first set of experiments, the adsorbed HI pool was incubated with 8 µg of DnaK, or buffer alone, in the presence or absence of ATP, for 8 min, and the amount of insulin and DnaK adsorbed on the beads after the incubation was determined. The material released in solution after the incubation was also analyzed by centrifugation, to separate soluble and aggregated proteins, and the amount of DnaK associated with the aggregates was also quantified. In the absence of DnaK, 35% of the initially adsorbed HI detached from the bead surfaces (7 µg/20 µg), corresponding to the spontaneous detachment of HI aggregates from the surface. In the presence of DnaK, larger amounts of protein, which contained both DnaK and HI, were released from the surface (15.4 µg + 2.4 µg = 17.8 µg). Under these conditions, 77% of the initially adsorbed HI detached from the beads (15.4 µg/20 µg). This material could be recovered by centrifugation, showing that it contained only HI aggregates. Moreover, an ELISA confirmed that <0.1 µg of HI could be recovered in soluble form. No protein renaturation had thus taken place. A large fraction of DnaK (2.4 µg of 8 µg) cosedimented with the released HI material, and small but significant amounts of DnaK (0.2 µg) remained at the bead surface. This strongly suggested that
<figure><img src="image_61.png" /></figure>

**Figure 4.** DnaK prevents the formation of prefibrillar insulin aggregates on hydrophobic surfaces. (A) Time course of adsorption of HI on hydrophobic surfaces in the presence of 0.3 µM DnaK without ATP (●) compared to the adsorption in the absence of DnaK (○). (B) Time course of amyloid fiber formation on hydrophobic surfaces in the presence of 0.3 µM DnaK without ATP (●) compared to the aggregation in the absence of DnaK (○). (C and D) Two-stage experiments. (C) A HI solution was incubated in the presence (filled symbols, b–d) or absence (○, a) of 0.3 µM DnaK. At the indicated time, 1 mM ATP was added to the DnaK-containing sample (b and c) and the HI solution was further incubated. Bold lines represent the time during which DnaK protects HI from aggregation. HI aggregation was monitored by turbidity. (D) DnaK (0.3 µM) was added at the indicated time (Δt) after the beginning of HI agitation in a hydrophobic microplate, and the incubation was further continued for 8 – Δt hours (●). In comparison, the kinetics of HI aggregation as measured in panel C (a) is shown (○). HI aggregation was monitored by turbidity after 8 h.

<figure><img src="image_62.png" /></figure>

**Figure 5.** Stoichiometry of the interaction of DnaK with insulin adsorbed on hydrophobic surfaces. A HI solution was agitated in hydrophobic microplates for the indicated preincubation time. DnaK was then added at different concentrations, and the HI solution was further incubated for 18 h. The extent of HI aggregation was determined by ThT staining. A 2-fold increase in the level of ThT staining over background was used as a criterion for the onset of HI aggregation. The minimal amount of DnaK needed to prevent the onset of HI aggregation is represented as a function of preincubation time. The amount of adsorbed HI is also represented.

DnaK detached insulin aggregates from the bead surface, remaining associated to them. The time course of this process

was ~20 min (data not shown). In contrast, no DnaK could be detected on hydrophilic surfaces where similar amounts of HI had been preadsorbed (data not shown). Qualitatively similar results were obtained in the presence and absence of ATP. However, less insulin was detached, and ~50% of the DnaK remained bound to insulin aggregates in the presence of 1 mM ATP.

In a second set of experiments, DnaK (8 µg) was incubated with HI aggregates that had already been released in solution. Under these conditions, ~0.5 µg of DnaK was bound to 150 µg of a HI aggregate suspension. This shows that DnaK does not bind efficiently to HI aggregates already present in solution. DnaK therefore binds specifically to HI adsorbed on hydrophobic surfaces.

Making use of the affinity of DnaK for insulin adsorbed on hydrophobic surfaces, we subsequently designed an assay for the sensitive detection of aggregated insulin on material surfaces. Insulin (or BSA as a control) was incubated in the presence of hydrophobic or hydrophilic surfaces for 1 h. The adsorbed protein (1–2 µg) was incubated with increasing amounts of DnaK, in the presence or absence of DnaJ and/or ATP. After the sample had been gently washed, DnaK was detected using anti-DnaK antibodies in a chemoluminescence
<table><caption>Table 1. DnaK Binding to HI-Covered Surfaces and HI Aggregates and LVEALYL Aggregates in Solution<sup>a</sup></caption><thead><tr><th rowspan="3"></th><th colspan="3">fraction</th></tr><tr><th rowspan="2">adsorbed</th><th colspan="2">released from beads</th></tr><tr><th>aggregates</th><th>soluble</th></tr></thead><tbody><tr><td>Hydrophobic</td><td>Surfaces, No DnaK Added</td><td></td><td></td></tr><tr><td>human insulin (µg)</td><td>11 ± 2</td><td>7 ± 3</td><td>&lt;0.1</td></tr><tr><td>Hydrophobic</td><td>Surfaces, with DnaK (8 µg)</td><td></td><td></td></tr><tr><td>human insulin (µg)</td><td>4.6 ± 1</td><td>15.4 ± 2.5</td><td>&lt;0.1</td></tr><tr><td>DnaK (µg)</td><td>0.2 ± 0.1</td><td>2.4 ± 0.6</td><td>5.4 ± 0.7</td></tr><tr><td>DnaK:HI molar ratio</td><td>277</td><td>77</td><td></td></tr><tr><td colspan="4">Hydrophobic Surfaces, with DnaK (8 µg) and Aggregated Insulin in Solution</td></tr><tr><td>human insulin (µg)</td><td></td><td>150 ± 10</td><td></td></tr><tr><td>DnaK (µg)</td><td></td><td>0.5 ± 0.2</td><td></td></tr><tr><td>Molar ratio (HI:DnaK)</td><td></td><td>4200</td><td></td></tr><tr><td>Hydrophobic surfaces, with DnaK (1.5 µg)</td><td></td><td></td><td></td></tr><tr><td>LVEALYL (µg)</td><td></td><td>25</td><td></td></tr><tr><td>DnaK (µg)</td><td></td><td>0.5</td><td></td></tr><tr><td>Molar ratio (peptide:DnaK)</td><td></td><td>4000</td><td></td></tr></tbody></table>

<sup>a</sup>Acid-washed borosilicate glass beads (diameter of 1 mm) were siliconized by immersion in SurfaSil (Pierce, 10%, w/w) in acetone and stabilized by being cured at 100 °C for 1 h. The water contact angle was measured (DSA100 Krüss) (93.5 ± 3.5°). SurfaSil-treated beads were incubated with HI in TBS-M buffer overnight. The fully aggregated HI solution was removed, and the beads were washed three times with 500 µL of TBS-M. The initial amount of HI adsorbed on the beads was determined. DnaK was then added, in the presence or absence of ATP, and the beads were further incubated for 1 h at 37 °C under agitation. The total protein content and the amount of HI or DnaK were determined in three fractions: the one adsorbed on the beads, the aggregated one, and the soluble ones released from the beads and separated by centrifugation (5000g for 10 min). The total amount of protein was quantified by the BCA assay; the amount of soluble HI was quantified by an ELISA (I2018 mouse monoclonal anti-insulin antibody), and the amount of DnaK was determined using SDS-PAGE and Coomassie staining. Quantification was performed with ImageJ. For the interaction of DnaK with the aggregated LVEALYL peptide, the peptide solution was prepared at 1 mM glycine buffer (pH 2.5). This solution was incubated in hydrophobic 96-well plates overnight at 37 °C with agitation (1200 rpm). The resulting aggregates were centrifuged and washed three times in TBS-M buffer (pH 7.4) before being resuspended and incubated in TBS-M buffer containing 1.5 µg of DnaK over 30 min. The DnaK/peptide solution was then centrifuged and washed three times in TBS-M buffer before SDS-PAGE analysis.

assay (see Experimental Procedures). In Figure 6A, the amount of adsorbed DnaK is plotted as a function of the total amount of DnaK. In the absence of DnaJ, small but significant amounts of DnaK (26 ng) were bound to insulin adsorbed on hydrophobic surfaces compared to the amount of DnaK bound on adsorbed BSA (14 ng). In contrast, almost no DnaK was recovered when insulin was incubated with hydrophilic surfaces (data not shown). The amount of DnaK bound to adsorbed insulin was only slightly reduced in the presence of 1 mM ATP (23 ng). In the presence of DnaJ, much larger amounts of DnaK were bound to adsorbed HI (372 ng), but not on adsorbed BSA (27 ng). Addition of ATP reduced the amount of DnaK binding in the presence of DnaJ to 147 ng. To study the DnaJ requirement for DnaK binding, the amount of DnaK was kept constant at 2.5 µg (0.18 µM) and the DnaJ concentration was increased (Figure 6B). DnaK binding reached a maximum for amounts of DnaJ larger than 1.28 µg (0.16 µM), which corresponds to a 1:1 DnaJ:DnaK ratio. These

<figure><img src="image_63.png" /><figcaption>**Figure 6.** Amount of DnaK adsorbed on surface-bound HI. A 0.5 mg/mL insulin or 0.5 mg/mL BSA control solution (×) was agitated for 1 h in a hydrophobic microplate. After removal of the solution, a DnaK solution was added. (A) The DnaK solution was not supplemented with ATP (■), supplemented with 1 mM ATP (□), and supplemented with DnaJ (0.16 µM) in the presence of 1 mM ATP (○) or in the absence of ATP (●). The amount of adsorbed DnaK was measured using an ELISA as described in Experimental Procedures and is represented as a function of the total amount of DnaK added. (B) A 0.18 µM DnaK solution supplemented with different amounts of DnaJ was added to the adsorbed insulin (■) and the adsorbed BSA (□). The amount of adsorbed DnaK was measured using an ELISA as described in Experimental Procedures and is represented as a function of the total amount of DnaJ added. Lines are hand-drawn and provided as a guide for the eye.</figcaption></figure>

data show that DnaK specifically interacts with HI adsorbed on hydrophobic surfaces and that the molecular cochaperone DnaJ reinforces this binding. The weak binding of DnaK in the presence of ATP is consistent with its lack of a protective effect against aggregation.

**Competition between the LVEALYL Amyloidogenic Peptide and Bacterial Chaperones during HI Aggregation on Hydrophobic Surfaces.** Recent studies by Ivanova et al.<sup>13</sup> pinpointed the role of two HI amino acid stretches that are likely to change extensively their conformation when HI spontaneously goes from a soluble form to amyloid aggregates at pH 2 and 60 °C. These sequences are SLYQLENY (A12–19) and LVEALYL (B11–17). Both contain hydrophobic residues and are mainly involved in α-helices in the HI monomer. In amyloid fibers, two LVEALYL stretches from two monomers are assumed to associate into antiparallel extended β-sheets that tightly interact via their hydrophobic side chains. The SLYQLENY sequence shows similar properties and also contributes to the formation of fibrillar structures. In this model, the core of HI amyloid fibers is therefore provided by hydrophobic interactions between these polypeptides and stabilized by the stacking of extended β-sheets. To investigate whether hydrophobic surfaces drive similar conformational changes, we studied the effect of the LVEALYL and the SLYQLENY peptides on HI aggregation kinetics. The SLYQLENY peptide
had no effect on HI aggregation kinetics, whereas the LVEALYL peptide strongly reduced the lag time in HI aggregation kinetics at substoichiometric concentrations relative to HI (Figure 7A). At higher concentrations, the

<figure><img src="image_64.png" /><figcaption>**Figure 7.** Effect of the LVEALYL peptide on HI aggregation kinetics. (A) A HI solution was agitated in a hydrophobic microplate in the presence of different amounts of the LVEALYL peptide, and aggregation was monitored using ThT fluorescence. The lag time (as defined in Experimental Procedures) is represented as a function of peptide concentration. (B) Aggregation kinetics of HI alone (O) or HI supplemented with 26 µM LVEALYL peptide (Δ), with 0.3 µM DnaK (●), or with both peptide and DnaK (▲). Aggregation was assessed using ThT fluorescence. Lines are hand-drawn and provided as a guide for the eye. (C) A HI solution was agitated in a hydrophobic microplate in the presence of the LVEALYL peptide (8 µM), DnaK (0.3 µM), and DnaJ (0.16 µM), as indicated. Aggregation was monitored using ThT fluorescence, and the lag time was determined. In the presence of DnaK, the lag time extended beyond 10 h.</figcaption></figure>

LVEALYL peptide inhibited the growth of HI amyloid aggregates, as reported previously at pH 2.¹³ The effect of the LVEALYL peptide was observed only in the presence of hydrophobic surfaces, the HI solution remaining perfectly stable in hydrophilic multiwell plates, whatever the peptide concentration. It should be noted that none of the peptides alone aggregate in hydrophobic or hydrophilic plates under the same pH, temperature, and concentration conditions.

Rüdiger et al. showed that the SHLVEALYLVCGER and CTSICSLYQLENYCN sequences in insulin bind DnaK.¹² Both encompass the aforementioned peptides involved in amyloid fiber formation (underlined). The same portions of insulin sequence are likely to be recognized by DnaJ, because it shares its substrates with DnaK.⁵ It follows that DnaK should antagonize the aggregation promoting effects of the LVEALYL peptide. As shown in Figure 7B, adding 26 µM LVEALYL peptide reduced the HI aggregation lag time from 2.5 ± 0.2 to 1.1 ± 0.2 h. The addition of 0.3 µM DnaK antagonized the effect of the LVEALYL peptide, extending the HI aggregation lag time up to 4.2 ± 0.7 h and slowing aggregation (Figure 7B). The presence of DnaJ enhanced the effect of DnaK [lag time of 7.8 ± 0.9 h (Figure 7C)]. More DnaK was needed to block HI aggregation for longer times. Conversely, ~60 µM LVEALYL peptide was needed to reduce the lag time to the value obtained in the absence of DnaK. Moreover, DnaK binds to amyloid aggregates formed by the LVEALYL peptide alone. Table 1 shows that 0.5 µg of DnaK binds to 25 µg of LVEALYL peptide aggregates.

## DISCUSSION

**DnaK Recognizes a Conformational Change Occurring on HI Adsorbed on Hydrophobic Surfaces.** During aggregation, three pools of insulin were evidenced in this study: the soluble insulin pool, the final amyloid aggregates, and insulin adsorbed on hydrophobic surfaces. In another article, we show that the latter pool contains prefibrillar insulin aggregates that are essential intermediates in the pathway(s) leading to insulin aggregation. The amount of DnaK needed to block insulin aggregation supports the view that DnaK binds to a minor portion of insulin present in the adsorbed insulin pool. The DnaK concentration (0.3–3 µM) is indeed much lower than the HI concentration in solution. Thus, DnaK cannot significantly displace the monomer–dimer–hexamer equilibria and therefore cannot significantly reduce the HI monomer concentration, which is the form of HI in solution that aggregates in the presence of hydrophobic surfaces.⁹ Because DnaK weakly interacts with preformed amyloid aggregates released in solution, the binding of DnaK to the end product of the aggregation reaction cannot explain its inhibitory effect (Table 1). The target of DnaK is therefore to be found at the material surface. Because the amount of DnaK needed to block HI aggregation increases during the lag time (Figure 5), we rule out competitive adsorption at the material surface as the reason for DnaK inhibition. Moreover, we show that DnaK directly binds to HI adsorbed on hydrophobic surfaces and to HI aggregates released from the surface (Figure 6 and Table 1). We therefore propose that DnaK blocks the formation of HI amyloid fibers on hydrophobic surfaces, by selective binding to target sequences in adsorbed HI. One of these targets is likely to be the hydrophobic LVEALYL peptide, aggregates of which also bind DnaK (see Table 1 and below).

There is kinetic competition between amyloid fiber growth and protection by DnaK, which is illustrated well by the effect of ATP and DnaJ. ATP-loaded DnaK is known to have a lower affinity for its protein substrate than ADP-loaded DnaK⁵ and DnaJ, which stimulates ATP hydrolysis on DnaK and reinforces its interaction with its substrate. Our results suggest that in the absence of nucleotides, DnaK binds strongly to adsorbed HI and DnaJ further stabilizes DnaK binding. When the mean residence time of DnaK on exposed hydrophobic HI stretches is sufficiently long, the formation of amyloid fibers is inhibited
(Figure 4). On the other hand, when the residence time of DnaK is reduced by addition of ATP, aggregates can form rapidly because of the liberation of previously occupied hydrophobic growing fiber ends (Figures 3 and 4). Thus, the absence of ATP, through a substrate affinity increase, allows DnaK to win the competition between adding a blocking DnaK to a growing fiber end and an additional HI molecule, which would promote fiber growth. The competition can also be promoted by a sufficiently high concentration of the LVEALYL peptide, which accelerates HI aggregation so that it overcomes DnaK protection (Figure 7). Although significant amounts of DnaK are released from the surface (Table 1), together with the HI amyloid aggregates, it is the surface-bound DnaK fraction that sets the pace for fiber growth. In the ATP-free state, this latter DnaK fraction exhibits an affinity for growing fiber ends that is sufficiently high to block fiber formation. The inhibitory effect of DnaK on HI aggregation is therefore linked to its presence on the material surfaces.

**The Amyloidogenic LVEALYL Peptide Is Exposed on HI during Its Aggregation on Hydrophobic Surfaces.**

The results of this work and others show that insulin aggregates are able to form and grow on hydrophobic surfaces. What is the likely mechanism of this self-assembly process? Up to 40 µg of HI accumulates on the 2 cm² surface area of a single microwell during aggregation (Figure 1A). This high surface concentration (0.2 g/m²) corresponds to 50 protein layers, assuming a uniform coverage of the surface. A consequence of this observation is that insulin does not merely adsorb on the plain hydrophobic surface, but on insulin already bound to the surface. The effect of the hydrophobic surface should therefore be transmitted from the bottom to the top HI layers, where incoming HI binds. This implies the existence of global conformational changes that relay the effect of the hydrophobic surface. Adsorption of HI molecules on a hydrophobic surface will itself trigger the exposure of a hydrophobic stretch at the surface of adsorbed HI. Although we did not evidence these conformational changes directly, several results support this hypothesis. (i) DnaK binds to HI adsorbed on hydrophobic surfaces, which shows that the structure of the HI protein has changed, exposing one of the two hydrophobic peptides known to bind DnaK: SLYQLENY and LVEALYL. (ii) The LVEALYL peptide itself interacts with HI in the presence of hydrophobic surfaces at a substoichiometric concentration in such a way to accelerate aggregation. This peptide has been shown to form antiparallel β-sheets with itself¹³ and could therefore help stabilize a conformational change in insulin by binding to intermediate states. (iii) The antagonist effects of the LVEALYL peptide and DnaK binding on HI aggregation reveal the presence of the same aggregation intermediates in the adsorbed HI pool. (iv) DnaK binds to preformed LVEALYL peptide aggregates. We therefore conclude that the buildup of amyloid fibers on the hydrophobic surface is due to HI adsorption and consequent conformational changes, exposing hydrophobic aggregation-prone sites in the protein. This situation occurs for some proteins, but not all of them. BSA, for instance, strongly binds to hydrophobic surfaces but does not accumulate on them and does not aggregate within an entire week in the presence of hydrophobic surfaces.

**Perspectives.** Our results have interesting potential biochemical and pharmaceutical applications. First, substoichiometric DnaK and DnaJ, in the absence of ATP, could represent a novel and convenient stabilizing additive. Indeed, the low DnaK concentration (0.3 µM), needed to stabilize HI solutions,

and its selectivity for exposed hydrophobic peptide domains are characteristics that most traditional stabilizing agents lack. Two recent papers by Rasmussen et al.³¹,³² showed that the protein α-crystallin, a member of the small heat shock protein family (sHSP), prevents HI aggregation on hydrophobic surfaces. Molecular chaperones, therefore, represent alternative approaches to guaranteeing the long-term storage of proteins, either in solution or in contact with the container surface. One should nevertheless be aware that DnaK triggers inflammatory responses and thus cannot be injected into patients.³³ DnaK, or better the DnaK-DnaJ combination, could also provide a sensitive in vitro assay to test for protein conformational changes at material surfaces, as shown for HI in this study. By revealing the presence of hydrophobic stretches exposed at the surface of proteins adsorbed on hydrophobic materials, this assay could provide a screening tool for the optimization of protein stability conditions.

## AUTHOR INFORMATION

### Corresponding Author

*Telephone: ++ 33 4 56 52 93 35. Fax: ++ 33 4 56 52 93 01. E-mail: marianne.weidenhaupt@grenoble-inp.fr.

### Present Address

§EIFFAGE Travaux Publics, Centre d'Etudes et de Recherches, 8 rue du Dauphiné, 69960 Corbas, France.

### Funding

This project was financed by a CNRS "Prise de risques" grant (CHAPROMAT). T.B. is a recipient of a CIFRE fellowship (371/2007). L.N.holds a doctoral fellowship from La Région Rhône-Alpes.

### Notes

The authors declare no competing financial interest.

## ACKNOWLEDGMENTS

We thank Benoît Duroux, Frédérique Crozet, and Sébastien Janvier for excellent technical assistance in particle size analysis, Laetitia Rapenne and Isabelle Paintrand in EM analysis and Bernd Bukau, Alexander Mogk, and Matthias Mayer for providing chaperone plasmids and protocols. We are grateful for the scientific input of Yves Bréchet (Grenoble-INP). We thank the CIME facility and BBSI laboratory (CEA Grenoble) for access to their equipment.

## REFERENCES

(1) Ellis, R. J., and van der Vies, S. M. (1991) Molecular chaperones. *Annu. Rev. Biochem.* **60**, 321–347.

(2) Ellis, R. J., van der Vies, S. M., and Hemmingsen, S. M. (1989) The molecular chaperone concept. *Biochem. Soc. Symp.* **55**, 145–153.

(3) Ellis, R. J., and Hemmingsen, S. M. (1989) Molecular chaperones: Proteins essential for the biogenesis of some macro-molecular structures. *Trends Biochem. Sci.* **14**, 339–342.

(4) Mogk, A., Tomoyasu, T., Goloubinoff, P., Rudiger, S., Roder, D., Langen, H., and Bukau, B. (1999) Identification of thermolabile *Escherichia coli* proteins: Prevention and reversion of aggregation by DnaK and ClpB. *EMBO J.* **18**, 6934–6949.

(5) Bukau, B., and Horwich, A. L. (1998) The Hsp70 and Hsp60 chaperone machines. *Cell* **92**, 351–366.

(6) Goloubinoff, P., Mogk, A., Zvi, A. P., Tomoyasu, T., and Bukau, B. (1999) Sequential mechanism of solubilization and refolding of stable protein aggregates by a bichaperone network. *Proc. Natl. Acad. Sci. U.S.A.* **96**, 13732–13737.

(7) Diamant, S., Ben-Zvi, A. P., Bukau, B., and Goloubinoff, P. (2000) Size-dependent disaggregation of stable protein aggregates by the DnaK chaperone machinery. *J. Biol. Chem.* **275**, 21107–21113.
(8) Sluzky, V., Tamada, J. A., Klibanov, A. M., and Langer, R. (1991) Kinetics of insulin aggregation in aqueous solutions upon agitation in the presence of hydrophobic surfaces. *Proc. Natl. Acad. Sci. U.S.A.* 88, 9377–9381.

(9) Sluzky, V., Klibanov, A. M., and Langer, R. (1992) Mechanism of insulin aggregation and stabilization in agitated aqueous solutions. *Biotechnol. Bioeng.* 40, 895–903.

(10) Brange, J., Andersen, L., Laursen, E. D., Meyn, G., and Rasmussen, E. (1997) Toward understanding insulin fibrillation. *J. Pharm. Sci.* 86, 517–525.

(11) Sharp, J. S., Forrest, J. A., and Jones, R. A. (2002) Surface denaturation and amyloid fibril formation of insulin at model lipid-water interfaces. *Biochemistry* 41, 15810–15819.

(12) Rüdiger, S., Germeroth, L., Schneider-Mergener, J., and Bukau, B. (1997) Substrate specificity of the DnaK chaperone determined by screening cellulose-bound peptide libraries, *EMBO J* 16, 1507–582.

(13) Ivanova, M. I., Sievers, S. A., Sawaya, M. R., Wall, J. S., and Eisenberg, D. (2009) Molecular basis for insulin fibril assembly. *Proc. Natl. Acad. Sci. U.S.A.* 106, 18990–18995.

(14) Cegielska, A., and Georgopoulos, C. (1989) Functional domains of the *Escherichia coli* dnaK heat shock protein as revealed by mutational analysis. *J. Biol. Chem.* 264, 21122–21130.

(15) McCarty, J. S., and Walker, G. C. (1991) DnaK as a thermometer: Threonine-199 is site of autophosphorylation and is critical for ATPase activity. *Proc. Natl. Acad. Sci. U.S.A.* 88, 9513–9517.

(16) Buchberger, A., Schroder, H., Buttner, M., Valencia, A., and Bukau, B. (1994) A conserved loop in the ATPase domain of the DnaK chaperone is essential for stable binding of GrpE. *Nat. Struct. Biol.* 1, 95–101.

(17) Veinger, L., Diamant, S., Buchner, J., and Goloubinoff, P. (1998) The small heat-shock protein IbpB from *Escherichia coli* stabilizes stress-denatured proteins for subsequent refolding by a multi-chaperone network. *J. Biol. Chem.* 273, 11032–11037.

(18) Olson, B. J., and Markwell, J. (2007) Assays for determination of protein concentration. *Current Protocols in Protein Science*, Chapter 3, Unit 3, 4, Wiley, New York.

(19) Smith, D. A., and Radford, S. E. (2000) Protein folding: Pulling back the frontiers. *Curr. Biol.* 10, R662–R664.

(20) Wiechelman, K. J., Braun, R. D., and Fitzpatrick, J. D. (1988) Investigation of the bicinchoninic acid protein assay: Identification of the groups responsible for color formation. *Anal. Biochem.* 175, 231–237.

(21) Stoscheck, C. M. (1990) Quantitation of protein. *Methods Enzymol.* 182, 50–68.

(22) LeVine, H. III (1999) Quantification of β-sheet amyloid fibril structures with thioflavin T. *Methods Enzymol.* 309, 274–284.

(23) Blundell, T. L., Cutfield, J. F., Dodson, G. G., Dodson, E., Hodgkin, D. C., and Mercola, D. (1971) The structure and biology of insulin. *Biochem. J.* 125, 50P–51P.

(24) Burke, M. J., and Rougvie, M. A. (1972) Cross-β protein structures. I. Insulin fibrils. *Biochemistry* 11, 2435–2439.

(25) Garriques, L. N., Frokjaer, S., Carpenter, J. F., and Brange, J. (2002) The effect of mutations on the structure of insulin fibrils studied by Fourier transform infrared (FTIR) spectroscopy and electron microscopy. *J. Pharm. Sci.* 91, 2473–2480.

(26) Theyssen, H., Schuster, H. P., Packschies, L., Bukau, B., and Reinstein, J. (1996) The second step of ATP binding to DnaK induces peptide release. *J. Mol. Biol.* 263, 657–670.

(27) McCarty, J. S., Buchberger, A., Reinstein, J., and Bukau, B. (1995) The role of ATP in the functional cycle of the DnaK chaperone system. *J. Mol. Biol.* 249, 126–137.

(28) Skowyra, D., and Wickner, S. (1995) GrpE Alters the Affinity of DnaK for ATP and Mg. *J. Biol. Chem.* 270, 26282–26285.

(29) Suh, W.-C., Burkholder, W. F., Lu, C. Z., Zhao, X., Gottesman, M. E., and Gross, C. A. (1998) Interaction of the Hsp70 molecular chaperone, DnaK, with its cochaperone DnaJ. *Proc. Natl. Acad. Sci. U.S.A.* 95, 15223–15228.

(30) Liberek, K., Wall, D., and Georgopoulos, C. (1995) The DnaJ chaperone catalytically activates the DnaK chaperone to preferentially bind the sigma 32 heat shock transcriptional regulator. *Proc. Natl. Acad. Sci. U.S.A.* 92, 6224–6228.

(31) Rasmussen, T., Kasimova, M. R., Jiskoot, W., and van de Weert, M. (2009) The Chaperone-like Protein Crystallin Dissociates Insulin Dimers and Hexamers, *Biochemistry* 4893139320.

(32) Rasmussen, T., Tantipolphan, R., van de Weert, M. and Jiskoot, W. (2010) The molecular chaperone alpha-crystallin as an excipient in an insulin formulation. *Pharm Res* 2713371347.

(33) Nolan, A., Weiden, M. D., Hoshino, Y., and Gold, J. A. (2004) Cd40 but not CD154 knockout mice have reduced inflammatory response in polymicrobial sepsis: A potential role for *Escherichia coli* heat shock protein 70 in CD40-mediated inflammation in vivo. *Shock* 22, 538–542.
<figure><img src="image_65.png" /></figure>

<figure><img src="image_66.png" /></figure>

# Human insulin adsorption kinetics, conformational changes and amyloidal aggregate formation on hydrophobic surfaces

Laurent Nault<sup>a</sup>, Perry Guo<sup>a</sup>, Bhagyashree Jain<sup>a</sup>, Yves Bréchet<sup>b</sup>, Franz Bruckert<sup>a</sup>, Marianne Weidenhaupt<sup>a,*</sup>

## ARTICLE INFO

**Article history:**

Received 12 June 2012

Received in revised form 14 September 2012

Accepted 19 September 2012

Available online 25 September 2012

**Keywords:**

Protein adsorption on material surfaces

Protein aggregation

Insulin

Surface plasmon resonance imaging (SPRi)

Fourier transform infrared spectroscopy
(FTIR)

## ABSTRACT

The formation of insulin amyloidal aggregates on material surfaces is a well-known phenomenon with important pharmaceutical and medical implications. Using surface plasmon resonance imaging, we monitor insulin adsorption on model hydrophobic surfaces in real time. Insulin adsorbs in two phases: first, a very fast phase (less than 1 min), where a protein monolayer forms, followed by a slower one that can last for at least 1 h, where multilayered protein aggregates are present. The dissociation kinetics reveals the presence of two insulin populations that slowly interconvert: a rapidly dissociating pool and a pool of strongly bound insulin aggregates. After 1 h of contact between the protein solution and the surface, the adsorbed insulin has practically stopped dissociating from the surface. The conformation of adsorbed insulin is probed by attenuated total reflection-Fourier transform infrared spectroscopy. Characteristic shifts in the amide A and amide II' bands are associated with insulin adsorption. The amide I band is also distinct from that of soluble or aggregated insulin, and it slowly evolves in time. A 1708 cm⁻¹ peak is observed, which characterizes insulin adsorbed for times longer than 30 min. Finally, Thioflavin T, a marker of extended β-sheet structures present in amyloid fibers, binds to adsorbed insulin after 30–40 min. Altogether, these results reveal that the conformational change induced in insulin upon binding to hydrophobic surfaces allows further insulin binding from the solution. Adsorbed insulin is thus an intermediate along the α-to-β structural transition that results in the formation of amyloidal fibers on these material surfaces.

© 2012 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

## 1. Introduction

Amyloid fibers are protein aggregates stabilized by intermolecular β-sheets [1]. They are characterized by staining with Congo red and Thioflavin T (ThT), and by a specific cross β X-ray diffraction pattern [2]. Several proteins or polypeptides are able to form amyloids under both natural and artificial conditions [3,4], e.g. islet amyloid polypeptide, which is associated with diabetes II, or the Aβ (1-40) peptide, which fibrillates in Alzheimer's disease [5]. Insulin is a well-known hormone that regulates blood glucose concentration. Its structure in solution is essentially helical [6]. Daily insulin injections are used to treat type I and many type II diabetic patients. It has long been recognized that insulin solutions also form amyloid fibers in the presence of hydrophobic materials [7-9], especially in contact with peristaltic pump tubing [10]. Furthermore, the daily injection of insulin solutions at the same place

occasionally induces local amyloidogenesis, characterized by the presence of insulin amyloidal aggregates [11,12]. Similarly, the formation of Aβ (1-40) peptide amyloid fibers is also strongly influenced by material surfaces [13,14], such as Teflon [15], alkane-thiol self-assembled monolayers [16] and the air-water interface [17,18]. Understanding the role of material surfaces in protein instability is therefore useful.

Pioneering work by Sluzky et al. [19,20] shows that human insulin (HI) indeed aggregates on hydrophobic material surfaces, at pH 7 and 37 °C under agitation, and that aggregation proceeds in two phases: a nucleation phase or lag time, where no insulin aggregation is observed, followed by an aggregation phase, where most of the insulin aggregates. The length of both phases is influenced by the material hydrophobicity, its surface area compared to the fluid volume, the extent of agitation and the temperature. Ballet et al. [21] further investigated the aggregation mechanism, and showed that the aggregates form on the surface and are later released in the fluid phase. Using DnaK, a bacterial chaperone, as a marker of protein denaturation, they showed that adsorbed HI has a conformation different from soluble and aggregated HI.

<footnote><sup>a</sup>Laboratoire des Matériaux et du Génie Physique, Phelma-Minatec, 3 parvis Louis Néel, BP257, F-38016 Grenoble Cedex, France</footnote>

<footnote><sup>b</sup>Laboratoire de Science et Ingénierie des Matériaux et des Procédés, Phelma-Campus, 1130 rue de la Piscine, BP 75, F-38402 Saint-Martin d'Hères Cedex, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: marianne.weidenhaupt-blackledge@grenoble-inp.fr (M. Weidenhaupt).</footnote>
A limitation of these previous studies is that the relatively small amount of adsorbed HI could not be detected with sufficient sensitivity during the nucleation time, with prefibrillar aggregates forming on the material surface. The evidence for the insulin conformational change upon adsorption was also indirect. Finally, in the 96-well plate configuration used in these experiments, ThT binding on adsorbed proteins was detected only after the onset of the aggregation phase. More sensitive methods are therefore required to monitor the adsorption and the interaction of HI molecules on hydrophobic model surfaces in real time.

In this study, we combine three biophysical approaches to study HI adsorption and conformational changes on hydrophobic surfaces (Fig. 1): surface plasmon resonance is sensitive to refractive index changes close to the surface and is used to monitor adsorption kinetics with good temporal resolution; attenuated total reflection-Fourier transform infrared spectroscopy (ATR-FTIR) monitors infrared vibrational bands and is therefore sensitive to conformational changes in surface-adsorbed protein, especially the conversion of α-helices into β-sheets; and ThT binds to amyloidal aggregates and is a sensitive marker of the final insulin state. The common trait of these experiments is that the surface of interest is put into contact with a continuous flow of an HI solution, in contrast with previous experiments, where an HI solution was incubated in a container with a specific surface chemistry. In the configuration presented here, the soluble HI concentration is almost constant throughout the duration of the experiment. A time-resolved sequence of events can thus be defined and studied, from the start of HI adsorption to the formation of amyloidal aggregates on the surface.

## 2. Materials and methods

### 2.1. Chemicals

Unless otherwise stated, all chemicals were purchased from Sigma-Aldrich. Experiments were conducted in TBS (25 mM Tris-HCl, pH 7.4, 125 mM NaCl). HI (recombinant, expressed in yeast) solutions were prepared at 0.5 mg ml⁻¹ (86 µM). All solutions were filtered (0.22 µm, Millipore Stericup filter unit) before use. For the ATR-FTIR experiments, insulin solutions were prepared at 86 µM in phosphate-buffered saline (10 mM H₃PO₄, 2.7 mM KCl and 137 mM NaCl) in D₂O and filtered at 0.22 µm. The pD was adjusted to 6.89 using a pH-meter, which corresponds to pH 7.30 in H₂O. The refractive index of the solutions was measured with an Abbe refractometer.

<figure><img src="image_67.png" /><figcaption>**Fig. 1.** Scheme of the experimental procedures used in this study. The sensor surface (gray bar) is coated with a hydrophobic self-assembled monolayer and a protein solution is passed over it. Contributions to the signal originate from the adsorbed protein layer and from the soluble protein contained in the penetration depth, which is technique dependent. The different physico-chemical parameters monitored during the studies are summarized below.</figcaption></figure>

### 2.2. Surface functionalization

Gold-coated prisms (SPRi Biochips) were obtained from Horiba. Prisms were cleaned using oxygen plasma (12 W, 2 min), and half of the gold surface was functionalized with polyethylene glycol thiol (SH-PEG, 8 monomer units, MW = 569.8 g mol⁻¹, from Iris Biotech GmbH) at 1 mg ml⁻¹ in water at room temperature to obtain a hydrophilic surface. The other half was then treated with hexadecane thiol (SH-C₁₆) at 1 mg ml⁻¹, diluted in toluene at room temperature to obtain a hydrophobic surface. The water surface contact angle was measured (DSA100 Krüss) and found to be 38 ± 3° and 100 ± 2.5° for the hydrophilic and hydrophobic sides, respectively. After each experiment, the prisms were washed with 1% sodium dodecyl sulfate (SDS) and distilled water to remove insulin aggregates.

Round (35 mm diameter) glass coverslips were cleaned using oxygen plasma (12 W, 2 min), siliconized with dimethyldichlorosilane (DDS) (5% in toluene, 2 h, room temperature), washed in toluene and ethanol, and cured at 110 °C for 2 h. The water surface contact angle was 102.1 ± 1.3°.

For ATR-FTIR measurements, a silicon prism was purchased from Specac Ltd, cleaned using oxygen plasma (12 W, 2 min), functionalized with phenyldimethylmethoxysilane (PDMMS) (5% in toluene, 2 h, room temperature), washed in toluene and ethanol, and cured at 110 °C for 2 h. The water surface contact angle was 103.1 ± 3.7°.

### 2.3. Surface plasmon resonance sensorgram acquisition and analysis

Buffer and protein solutions were injected in the SPRi chamber using syringe pumps at 100 µl min⁻¹ at room temperature (22 °C). Reflectivity images were taken every 3 s. The SPRi instrument (Horiba Scientific SPRi-Lab+™) records reflectivity changes of manually predefined spots on the treated gold surface. Calibration was performed using a 50 mM NaCl solution, to adjust the response of the spots to a known refractive index change (3 × 10⁻⁴). A background image was acquired at the beginning of the experiment, which was then subtracted from the subsequent ones. Three zones of interest were defined on the SH-PEG and SH-C₁₆ sides of the prism, and the sensorgrams obtained on each side were averaged. A difference sensorgram ΔR(t) was finally obtained by subtracting the SH-PEG sensorgram from the SH-C₁₆ one.

The adsorption kinetics were analyzed with the following equation:

$$ \Delta R(t) = A(1 - \exp(-kt)) + bt \quad (1) $$

where A and k are the amplitude and rate of the fast adsorption component, and b is the rate of the slow protein binding component.

The desorption kinetics were analyzed with the following equation:

$$ \Delta R(t) = A_1 \exp(-k_1 t) + A_2 \exp(-k_2 t) \quad (2) $$

where A₁ and A₂ are the amplitude of the fast and slow desorption components, respectively, and k₁ and k₂ are the rate constants of the fast and slow desorption components, respectively. Data are obtained from several independent experiments, using different functionalized prisms and HI solutions.

The conversion between the ΔR and the adsorbed mass per area is given in pg mm⁻² by the equation:

$$ \text{mass} = 122.25\Delta R - 0.6557 \quad (3) $$

assuming that the signal is in the linear range of the instrument and solution refractive index changes have been corrected (equation provided by Horiba SPRi+). The relationship between ΔR and the refractive index changes, Δn, was determined using NaCl solutions
at different concentrations and was found to be: ΔR = 28333Δn. This relation was used to calculate the amplitude of the refractive index change in protein solutions compared to buffer.

Having recorded the dissociation kinetics, the hydrophobic surface was regenerated with SDS (5%, 5 min).

## 2.4. FTIR recordings

Aggregated HI solutions were prepared in 96-well plates (Greiner Bio-One, contact angle = 85 ± 4.7°) as follows. The plates were covered by plastic sheets, incubated at 37 °C and shaken at 1200 rpm (Heidolph Titramax, 1.5 mm vibration orbit) for 12 h. The aggregated solution was analyzed using a Brucker Vertex 70 FTIR instrument. The liquid cell was a CaF₂ cell from Perkin Elmer Instruments. Using the OPUS software, compensation was applied to filter the contributions of H₂O and CO₂ to the spectra. Smoothing was done using the Savitzky-Golay equation. Baselines were flattened using a linear correction.

## 2.5. ATR-FTIR recordings

Buffer and protein solutions were prepared in D₂O and injected in the chamber using a peristaltic pump at 10 ml min⁻¹ in a closed circuit, at room temperature. A tubing diameter of 5 mm was used to reduce shear stress. No amyloid fibers were formed on the tubing during the time of the experiment. The total volume of the sample was 10 ml. During recording of the spectra (about 2 min), the flow was stopped. Spectra were acquired from 1400 to 4000 cm⁻¹. Baselines were flattened using a linear correction in the range 4000–2800 cm⁻¹ and in the range 2000–1400 cm⁻¹. H₂O and CO₂ compensations were done as described for FTIR recordings. Further smoothing was performed using a Savitzky-Golay algorithm. Spectral decomposition of the amide I band (1600–1700 cm⁻¹) was done using the second derivative of the spectra to find peak positions. Then, using the Levenberg-Marquardt algorithm, 3–5 peaks were fitted to the experimental data (residual relative error less than 2.5 × 10⁻⁵).

The spectra A(t) recorded at time t contain two contributions: A(t) = Aads(t) + Asol, where Aads(t) is the spectrum of adsorbed HI at time t and Asol is the constant soluble HI contribution. With this notation, the spectrum recorded after 80 min in the presence of HI and 1 min wash with buffer is Aads(80). The spectra of HI adsorbed at time t, Aads(t), can therefore be calculated by the following equation:

$$ 
\begin{aligned}
A_{ads}(t) &= A(t) - A(0) + A_{ads}(80) - [A(80) - A(0)] \\
&= A(t) + A_{ads}(80) - A(80)
\end{aligned}
\quad (4) $$

## 2.6. Surface fluorescence measurements

A flow chamber was built on a DDS-coated glass coverslip using a Glycotech flow chamber (channel width = 2.5 mm, channel thickness = 0.127 mm). A 20 µl min⁻¹ flow rate was applied by a syringe pump. When indicated, ThT was added to the buffer and protein solutions at a 20 µM final concentration. The surface fluorescence was observed by a ×63 objective (NA = 1.4) on an IX-71 Olympus microscope fitted with a DAPI fluorescence cube (λex = 435 ± 10 nm, λem = 485 ± 25 nm) and connected to an Olympus DP30BW camera. The image of the field stop was used to ensure proper focusing on the glass surface. Fluorescence images of the surface were recorded at the indicated times. To avoid photobleaching, each image was recorded at a different position, using a motorized stage. Using the ImageProPlus software, the average fluorescence

intensity was calculated on the image of the field stop and the value of the dark background was subtracted.

# 3. Results

## 3.1. Insulin adsorption and dissociation kinetics on hydrophobic surfaces

Surface plasmon resonance imaging (SPRi) is a sensitive method to measure the amount of molecules adsorbed on functionalized gold surfaces (see Fig. 1). A gold prism was prepared with half of the surface coated with SH-PEG and the other half with C₁₆ alkyl-thiol. SPR is sensitive to refractive index changes within the penetration distance of the evanescent wave. When a solution is bathing the prism, the SPR signal sums up two contributions, one due to the refractive index of the bulk solution, the other due to the mass adsorbed on the gold surface. The response time of the setup was determined by passing a 50 mM NaCl solution over the functionalized prism. In this case, no adsorption is expected. The sensorgrams indeed indicate a fully reversible, fast (<1 min) signal, the amplitude of which minimally depends on the surface functionalization (Fig. 2A and B). This corresponds to the time needed to exchange solutions over the prism surface. The linearity of the technique with respect to the amount of adsorbed material was assessed by adsorbing a polyelectrolyte multilayer film on the prism layer by layer. In this experiment, the prism was not functionalized and a 10 mg ml⁻¹ polyethyleneimine (PEI) solution in TBS was passed first to provide an anchoring layer. Solutions of 5 mg ml⁻¹ polystyrene sulfonate (PSS, negatively charged) and 5 mg ml⁻¹ polyallylaminehydrochloride (PAH, positively charged) in TBS were passed alternately over the surface, separated by washes with TBS (Fig. 2C). Previous work showed that the thickness of these films increased linearly with the number of layers deposited, each polystyrene sulfonate layer forming a 3–5 nm thick layer [22]. Using QCM-D and waveguide spectroscopy, up to 12 layers could be monitored. In contrast, with SPRi, the deposition of the same polyelectrolytes gave signals that rapidly decreased as the number of layers increased (Fig. 2C and D). The amplitude of the signal was plotted as a function of the thickness of the multilayered film, obtained by waveguide spectroscopy [23] and fitted as S[1 − exp(−z/dₚ)], where S represents the maximum reflectivity change and dₚ is the penetration distance. From Fig. 2C and D, a penetration distance of 18.7 ± 1 nm and a maximum signal of 26.5 ± 1 RU can be estimated. A single layer of PSS corresponds to 6 RU, which is 0.73 mg m⁻² according to Eq. (3).

A typical HI adsorption kinetics experiment is shown in Fig. 3A. A 86 µM HI solution (0.5 mg ml⁻¹) was passed over the functionalized prism during 14 min, then the surface was washed with buffer for 45 min. Sensorgrams were recorded on both the hydrophilic SH-PEG-coated side and the hydrophobic C₁₆ alkyl-thiol-coated side. On the hydrophilic side, a fast reversible signal was observed, which could be attributed to refractive index changes solely. On the hydrophobic side, a much larger signal was observed, which rose in two phases and decreased much more slowly after the buffer had replaced the protein solution. This signal is due to both the refractive index change and the binding of HI on the hydrophobic surface. After a 5 min wash, the reflectivity remained 0.5 RU higher than on the PEG-coated side.

Subtracting the sensorgram acquired on the PEG-coated surface from the one acquired on the C₁₆ alkyl-coated surface allows the common component due to refractive index changes to be eliminated and the HI adsorption and desorption kinetics on the hydrophobic surface to be specifically obtained (Fig. 3B). The association curve consists of two components – a fast signal, rising to 1 RU in
<figure><img src="image_68.png" /><figcaption>**Fig. 2.** Calibration of the SPRi apparatus. A and B. Changes in C₁₆-coated (heavy line) and PEG-coated (thin line) gold surface reflectivity were monitored by surface plasmon resonance during injection of 50 mM NaCl (A) and washing in buffer (B). (C) SPRi recording of a PSS and PAH polyelectrolyte layer-by-layer film built on a gold surface. PEI is first deposited as an anchoring layer. (D) The amplitude of the SPRi signal after depositing the successive layers and washing with buffer (data from C) is plotted as a function of the adsorbed layer thickness (z) determined by Picart et al. [23]. The solid line is a fit with the equation: $S(z) = S[1 - \exp(-z/d_p)]$, where $S$ (26.5 ± 1 RU) is the maximum reflectivity change and $d_p$ (18.7 ± 1 nm) is the penetration distance.</figcaption></figure>

about 1 min, followed by a slow, almost linear increase. During the wash, the dissociation curve also consists of two components – a fast signal, going up to 1.3 RU in about 5 min, followed by a much slower decrease that lasted for more than 30 min. The association and dissociation kinetics were adequately fitted by the phenomenological equations 1 and 2 (see Materials and methods), giving seven parameters: A, k, b, A₁, k₁, A₂ and k₂. A and k represent the amplitude and rate constant of the fast association component, and b represents the slope of the slow association component. A₁ and k₁ (and A₂ and k₂) represent the amplitudes and rate constants of the fast (and slow) dissociation components, respectively. These biphasic association and dissociation curves show that HI binding

<figure><img src="image_69.png" /><figcaption>**Fig. 3.** HI association and dissociation kinetics on hydrophobic surfaces. (A) SPRi signal for 0.5 mg ml⁻¹ HI injection and buffer washing on a prism functionalized with C₁₆-thiol (heavy line) or SH-PEG (thin line). (B) The difference signal obtained by subtracting the signals recorded on the SH-PEG and SH-C₁₆ sides reveals the HI adsorption and desorption kinetics (black line). The red and green curves represent a mathematical fit of the data with Eqs. (1) and (2), respectively. A = 1.076 ± 0.005; k = 1.32 ± 0.02; b = 0.0545 ± 0.0005. A₁ = 1.153 ± 0.005; k₁ = 0.707 ± 0.007; A₂ = 0.570 ± 0.004; k₂ = 0.0595 ± 0.0007.</figcaption></figure>

on hydrophobic surfaces is not a reversible process. The slow dissociation component indicates that some HI is strongly bound to the surface, whereas the fast dissociation component suggests that another pool of HI is much more weakly bound to the surface.

In a parallel experiment, the surface concentrations of HI adsorbed on a hydrophilic PEG-coated microwell or on a hydrophobic polystyrene plastic surface (2 cm²) were directly determined using the bicinchoninic assay [24]. No HI was detectable on the hydrophilic surface, whereas 0.4 mg m⁻² was quantified on hydrophobic surfaces. Comparing this value to the corresponding SPRi experiments shows that 1 RU = 0.2 mg m⁻². This is close to the value obtained with Eq. (3) (0.12 mg m⁻² per RU). Assuming that the calibration determined for the fast phase of the SPRi association signal holds also for the slow phase, the rate of insulin adsorption would be about 0.013 mg m⁻² min⁻¹.

Since SPRi is a very sensitive technique, HI adsorption could easily be monitored on hydrophobic surfaces at different concentrations down to 0.1 µM. Fig. 4A shows an SPRi difference sensorgram obtained on the same surface where successive HI concentrations ranging from 0.49 to 86 µM were applied. Each HI concentration was applied for 10 min, after which the surfaces were washed. It is apparent from the difference sensorgram that the slope of the slow association kinetics and the amplitude of the slow dissociation kinetics both increase with HI concentration. In Fig. 4B and C, the effect of increasing HI concentrations on the parameters A, k (Fig 4B) and b (Fig 4C) describing the association kinetics are displayed. The amplitude A of the fast association component increases with HI concentration and saturates at 1 ± 0.05 RU (2.6 mg m⁻²), with an apparent binding constant of 2.5 ± 0.5 µM, showing that the amount of HI was limited by the available surface. The rate constant of the fast association kinetics k remains constant at 1.3 ± 0.1 min⁻¹. The fast rise of the
<figure><img src="image_70.png" /><figcaption>**Fig. 4.** Effect of HI concentration on HI adsorption and desorption kinetics. (A) Successive HI adsorption and desorption difference SPRi signals recorded at the indicated HI concentrations. (B, C) The adsorption kinetics were fitted with Eq. (1) to calculate the amplitude *A* and the rate constant *k* of the fast component and the slope *b* of the slow component. (B) Parameters *A* and *k* are plotted as a function of the HI concentration. The error in *A* is smaller than the symbol size. The solid curve is a fit with the following relationship: $A = A_{max}[HI]/([HI] + K)$, where $A_{max}$ (1 ± 0.05 RU) is the HI binding site concentration and *K* (2.5 ± 0.5 µM) is the apparent binding constant. (C) Parameter *b* is plotted as a function of the HI concentration. The error on *b* is smaller than the symbol size. The solid curve is a fit with the following relationship: $b = b_{max}[HI]/([HI] + K')$, where $b_{max}$ (0.058 ± 0.002 RU min⁻¹) is the maximum HI association rate and *K'* (6.1 ± 1 µM) is the apparent binding constant.</figcaption></figure>

association kinetics is thus limited by the response time of the SPRi setup. The slope of the slow association component *b* also increases with HI concentration and saturates at 0.058 ± 0.002 RU min⁻¹ (0.023 ± 0.001 mg m⁻² min⁻¹), with an apparent binding constant of 6.1 ± 1 µM. The growth of HI aggregates on the hydrophobic surface thus occurs at a limited number of specific sites.

In a second set of experiments, we varied the time during which the HI solution was in contact with the prism surface (Fig 5A). As the contact time is increased, the slow association component lasts longer (not shown) and the slow dissociation component becomes more important (Fig 5B) and slower (Fig 5C). This indicates that, as time progresses, insulin binds more strongly to the alkyl surface and an increasing amount of insulin remains adsorbed in a strongly bound form on the hydrophobic surface. The parameters describing the association kinetics are similar to those described in the experiments where the HI concentration was varied. For the dissociation kinetics, both sets of experiments were analyzed together,

<figure><img src="image_71.png" /><figcaption>**Fig. 5.** Effect of incubation time on HI desorption kinetics. (A) SPRi desorption difference sensorgrams obtained after application of 86 µM HI for the indicated time. The SPRi difference signals have been normalized to the values reached at the beginning of the washing with buffer. (B) Amplitudes of the fast (open symbols) and slow (closed symbols) components of the desorption sensorgrams, represented as a function of the SPRi value at the onset of the washing with buffer. They correspond to parameters $A_1$ and $A_2$ in Eq. (2). The lines are hand-drawn. (C) Rate constant of the fast (open symbols) and slow (closed symbols) components of the desorption sensorgrams, represented as a function of the SPRi value at the onset of the buffer wash. They correspond to parameters $k_1$ and $k_2$ in Eq. (2). The lines are exponential fits of the data. In the case of the slow component, the two extreme points were omitted. In (B) and (C), squares and circles correspond to desorption kinetics obtained after applying either different HI concentrations for the same time or 86 µM HI for different times, respectively. The errors in the determination of the kinetic desorption parameters are smaller than the symbol size.</figcaption></figure>

and the kinetic parameters were plotted as a function of the amount of HI adsorbed on the surface at the beginning of the wash. The relative amplitude of the fast dissociation component $A_1$ decreased and the amplitude of the slow dissociation component $A_2$ increased as the amount of adsorbed HI increased. Furthermore, the fast and the slow dissociation rate constants $k_1$ and $k_2$ decreased exponentially, in a parallel manner, as the amount of adsorbed HI increased. This shows that the presence of adsorbed HI further stabilizes HI binding. HI adsorption is therefore a cooperative process.

It has been shown that agitation has two opposite effects on HI aggregation induced by hydrophobic surfaces: on the one hand, it prevents the formation of a depleted diffusion layer that slows
<figure><img src="image_72.png" /><figcaption>**Fig. 6.** ATR-FTIR spectra of HI adsorbed on hydrophobic surfaces. In (A), (B) and (C), the left and the right panels display the spectra in the 4000–2800 cm⁻¹ or the 2000–1400 cm⁻¹ ranges, respectively. In the left and right panels, the baseline was zeroed at 4000 or 2000 cm⁻¹, respectively, except in the right panel of (A), where the zero was set at the minimum. Data are representative of three experiments. (A) Transmission FTIR spectra of soluble (blue) and aggregated (red) HI. ATR-FTIR spectrum of HI adsorbed for 80 min on a hydrophobic surface and washed (green). The spectra have been normalized to the same total area. (B) Successive ATR-FTIR spectra recorded on the same hydrophobic surface while passing an 86 µM HI solution during 80 min. The time is indicated in the right panel. (C) Same data as in (B). The ATR-FTIR spectrum recorded after 1 min has been subtracted from those recorded later to show the increase in adsorbed HI. In addition, the spectrum obtained after 80 min of contact with the HI solution and a 1 min wash is also displayed. (D) Total absorbance of adsorbed HI calculated according to Eq. (3) integrated over the 4000–2800 cm⁻¹ and the 2000–1400 cm⁻¹ ranges, as a function of time. For comparison, the association SPRi difference signal is represented as a thick line. (E) Amide I band of soluble HI (blue), aggregated HI (red) and HI adsorbed for 80 min on hydrophobic surfaces (green). Data are from (A). The baseline is linearly adjusted between 1700 and 1570 cm⁻¹ for soluble HI and aggregated HI spectra, and between 1730 and 1570 cm⁻¹ for the adsorbed HI spectrum. In the case of the adsorbed HI spectrum, the decomposition of the band into four peaks is displayed as thin lines that represent the individual peaks and their sum.</figcaption></figure>

down the accumulation of protein on the surface; on the other hand, it helps to detach already formed aggregates. The fluid flow rate was therefore changed to mimic the effect of agitation (data not shown). The rate of HI adsorption was monitored as the fluid flow was stopped. No change was observed, showing that HI diffusion is fast enough to allow HI aggregation at surfaces.

### 3.2. Insulin conformational changes on hydrophobic surfaces

Insulin conformational changes during its adsorption on hydrophobic surfaces were investigated by ATR-FTIR (Fig. 1). The infrared spectrum of a protein is indeed sensitive to its secondary structure. For this purpose, the top contact surface of an ATR-FTIR
silicon prism was functionalized with phenyldimethylmethoxysilane, which forms a hydrophobic self-assembled monolayer. The baseline spectrum was recorded with the buffer solution. An 86 µM HI solution prepared in $^2$H$_2$O was then passed over the prism and 4000–1400 cm$^{-1}$ infrared spectra were recorded at the times indicated during 80 min at room temperature (20 °C). Since recording a single spectrum takes about 1 min, the infrared spectrum referring to time point 1 corresponds to insulin already adsorbed on the prism for 1–2 min. Infrared spectra were therefore recorded early after the fast initial HI adsorption (time 1) and during the slow phase of the HI adsorption kinetics (subsequent times).

In comparison, Fig. 6A shows the spectra of the initial HI solution and of a suspension of aggregated HI, obtained after agitation in a hydrophobic multi-well plate [21], both recorded by transmission FTIR. The spectra are displayed in two parts, the 4000–2800 cm$^{-1}$ region in the left panel and the 2000–1400 cm$^{-1}$ region in the right one, skipping the intermediate 2800–2000 cm$^{-1}$ region, where the signal is almost flat or does not contain any protein contributions. In proteins, the peptide bond (amide group) is the main contributor to the infrared spectrum, but specific amino acid side chains may also contribute significantly in spectral regions where no other chemical group absorbs, e.g. tyrosines at 1515 cm$^{-1}$. In the 4000–1400 cm$^{-1}$ range, amide groups contribute to three groups of peaks: the amide A peak, in the 3600–3100 cm$^{-1}$ range, the amide I peaks, in the 1700–1600 cm$^{-1}$ range, and the amide II' peaks, in the 1500–1400 cm$^{-1}$ range [25]. The prime refers to the fact that in the presence of $^2$H$_2$O, hydrogen is exchanged for deuterium in the amide NH group. The soluble and aggregated HI spectra are similar in the amide A/B and amide II' bands (3420, 3100 and 1460 cm$^{-1}$, respectively), but are characteristically different in the amide I band, with one prominent 1650 cm$^{-1}$ band for soluble HI and two bands around 1660 and 1630 cm$^{-1}$ for aggregated HI, as reported previously [26].

The successive ATR-FTIR spectra clearly show the progressive accumulation of protein material on the PDMMS surface (Fig. 6B). The first spectrum (time 1) is very similar to the spectrum of soluble HI and displays the amide A peak at 3420 cm$^{-1}$, the amide I peak at 1650 cm$^{-1}$, the tiny tyrosine contribution at 1515 cm$^{-1}$ and the amide II' peak at 1450 cm$^{-1}$. At later times, however, specific peaks appear and increase rapidly, especially at 3276 and 3204 cm$^{-1}$ in the amide A band and at 1705 cm$^{-1}$ in the amide I band. Since the penetration distance of the infrared evanescent waves in ATR-FTIR is about 2 µm, the recorded spectra are indeed the sum of the soluble HI contribution and the adsorbed HI contribution. Assuming that the adsorbed HI layer remains negligible compared to the 2 µm penetration depth, the contribution of soluble HI is constant during the association phase. Difference spectra thus reveal the adsorbed HI contribution. In Fig. 6C, the spectrum recorded at $t=0$ has been subtracted from subsequent spectra. These difference spectra are quite similar in shape, except in the range 1730–1670 cm$^{-1}$ (see below), but are clearly distinct from the spectrum of HI in solution or aggregated (Fig. 6A). Besides the aforementioned amide A and amide I differences, the amide II' is clearly shifted to 1435 cm$^{-1}$. Furthermore, these difference spectra are very similar to the ATR-FTIR spectrum recorded at the end of the experiment, once HI had been passed over the prism surface for 80 min and washed with buffer for 1 min (black curve in Fig. 6C). This proves that adsorbed HI has a specific spectrum and that spectra recorded subsequently in time (Fig. 6B) are the sum of a constant soluble HI contribution and increasing adsorbed HI contributions. Curve fitting confirmed this analysis, except in the 1730–1670 cm$^{-1}$ region (see below). After washing with 1% SDS, the flat initial baseline was restored, showing that this detergent is able to remove all adsorbed proteins. The spectrum recorded after buffer wash is therefore due to adsorbed HI. In Fig. 6A, this

spectrum has been normalized and displayed along the soluble and aggregated HI ones to facilitate their comparison.

The difference between the spectrum obtained after washing the prism surface (Fig. 6C, black curve) and the last difference spectrum (80 min – 1 min, Fig. 6C, pink curve) corresponds to the HI adsorbed during the first minute. It is therefore possible to calculate the spectrum of adsorbed HI as a function of time from Eq. (3). In Fig. 6D, the absorbance is summed over the 4000–2800 cm$^{-1}$ and 2000–1400 cm$^{-1}$ ranges, and plotted as a function of time. The fact that the integrated absorbance continuously increases over 80 min confirms that HI is indeed able to adsorb not only on the hydrophobic surface, but also on HI molecules already adsorbed on the surface. To compare this plot with the SPRi results, the SPRi association signal was also plotted, and displayed using a scale that allows its comparison to the integrated absorbance signal. The integrated absorbance signal is quite comparable to the SPRi signal: it rises rapidly during the first 10 min (fast component), then increases linearly for about 1 h (slow component). It should nevertheless be noted that the FTIR signal rises less rapidly than the SPRi signal in the first 10 min.

The amide I band of adsorbed HI is strikingly different from that of soluble and aggregated HI. First, the entire band is shifted from the 1700–1600 cm$^{-1}$ range to 1730–1630 cm$^{-1}$. Second, the substructure of the amide I band exhibits four main peaks, at 1635, 1655, 1687 and 1708 cm$^{-1}$ (Fig. 6E). The relative contributions of these peaks to the amide I band are 18, 33, 29 and 20%, respectively. A detailed analysis of the evolution of the amide I band of adsorbed HI with time reveals that, initially, the 1708 cm$^{-1}$ peak is minimal, but increases over about 30–40 min (Fig. 7A and B). In contrast, the 1655 cm$^{-1}$ peak reaches its maximum level in less than 5 min and remains almost constant thereafter. The evolution of the 1708 cm$^{-1}$ peak parallels that of the relative amplitude of the slow dissociation component of the SPRi signal (Fig. 7B). This documents the complexity of conformational changes and reveals the existence of slow conformational rearrangements within adsorbed HI.

### 3.3. Insulin aggregate formation on hydrophobic surfaces

Amyloidal aggregates are characterized by the formation of large intermolecular β-sheets, which can be probed by the binding of ThT, resulting in a specific fluorescence signal, distinct from soluble ThT fluorescence. We used a Glycotech flow chamber to pass an HI solution over a surface of interest in the presence of ThT and monitor the fluorescence associated with the surface. The coverslip was functionalized with dimethyldichlorosilane, which forms a hydrophobic self-assembled monolayer on glass. Fig. 8A shows a typical experiment. When a buffer solution containing ThT was introduced in the chamber, a small but significant surface fluorescence signal was detected. Washing the surface with buffer alone resulted in a fast decrease in surface fluorescence down to the background level, showing that the fluorescence signal was due to ThT that was loosely bound to the hydrophobic surface (Fig 8A). When an 86 µM HI solution containing ThT was passed over the coverslip, the fluorescence signal first decreased close to zero, then increased sharply after 40 min and was saturated by about 90 min. No ThT fluorescence signal appeared on a hydrophilic surface (plain glass coverslip, data not shown). Passing a BSA solution over the hydrophobic surface also decreased the surface fluorescence signal, but gave rise to little or no fluorescence recovery. We therefore interpret the surface fluorescence decrease as a strong competitive protein binding, replacing the weakly bound ThT. The fluorescence increase is due to the formation of insulin amyloid aggregates on the surface. BSA indeed does not readily form amyloid aggregates on hydrophobic surfaces (data not shown). Once HI had accumulated on the surface, washing with a
<figure><img src="image_73.png" /><figcaption>**Fig. 7.** (A) Evolution of the HI amide I band as a function of the time of contact between an 86 µM HI solution and the hydrophobic surface. Data are from Fig. 6. The adsorbed HI spectra are calculated using Eq. (3). The baselines of the amide I bands are linearly adjusted between 1730 and 1570 cm⁻¹. (B) The infrared absorbances of adsorbed HI at 1655 cm⁻¹ (closed circles) and at 1708 cm⁻¹ (open circles), and the relative amplitude of the slow component of the SPRi dissociation signal (squares), are plotted as a function of the time of contact between an 86 µM HI solution and the hydrophobic surface. The lines are hand-drawn.</figcaption></figure>

buffer solution devoid of ThT resulted in a very slow fluorescence decrease, showing that ThT was now strongly bound to HI adsorbed on the surface (Fig. 8B). Quantitatively, the ThT dissociation kinetics was five times lower from adsorbed HI than from plain hydrophobic surfaces. The surface fluorescence decreased very slowly (typically for 15 h) during washing with a buffer solution containing ThT, confirming that amyloid HI does not easily dissociate from the surface (Fig. 8B). In contrast, washing with SDS allowed a fast and complete disappearance of the surface fluorescence (data not shown). Altogether, these results show that a lag phase of 40 min precedes the appearance of ThT positive aggregates on hydrophobic surfaces, that ThT binds more strongly to adsorbed HI than to hydrophobic surfaces and that ThT-positive aggregates are strongly bound to the surface.

## 4. Discussion

In this work, we reconstituted the formation of insulin amyloid aggregates on hydrophobic surfaces and delineated in time a series of sequential molecular events that occur on the material surface (Fig. 9). Amyloid aggregates, evidenced by ThT fluorescence, appear about 40 min after the hydrophobic surface is put into contact with an insulin solution. It should be noted that the experimental conditions are slightly different from those used in previous insulin aggregation assays such as in Slutzky et al. [19,20] or Ballet et al. [21]. First, the protein solution flows over the surface of interest, instead of being simply incubated. The bulk protein concentration therefore remains constant, except during ATR-FTIR experiments, where the solution is recirculated. Note that in this case, the total HI content of the setup (5 mg) is in large excess to the HI expected

<figure><img src="image_74.png" /><figcaption>**Fig. 8.** Formation of HI amyloid fibers on hydrophobic surfaces. The surface fluorescence of a PDMMS-coated coverslip is monitored to detect the presence of bound ThT. The fluorescence levels are normalized to the value obtained after passing a 20 µM ThT solution for 120 min on the surface. The curves displayed are representative of three independent experiments. (A) At *t* = 0, either TBS buffer without ThT (black squares) or TBS buffer containing 0.5 mg ml⁻¹ (6.5 µM) BSA and 20 µM ThT (open triangle) or TBS buffer containing 0.5 mg ml⁻¹ (86 µM) HI and 20 µM ThT (open circle) are passed over the surface. Surface fluorescence was measured every 10 min. The lines are hand-drawn. (B) After passing a 20 µM ThT solution for 120 min on the surface, the surface is washed at *t* = 0 with either ThT-free TBS buffer (closed circles) or ThT-containing buffer (open circles). The lines represent a linear fit to the data.</figcaption></figure>

to bind to the tubing and the ATR-FTIR sensor area (4 cm²; after 1 h, the bound HI is $4 \times 10^{-4}$ cm² × (0.2 mg cm⁻² + 0.013 mg cm⁻² min⁻¹ × 60 min) = 3.9 µg, according to Eq. (1)). Second, the experimental temperature, a key parameter in protein unfolding, was 20 °C instead of 37 °C. From the temperature dependence studied by Ballet et al. [24], the lag time before the onset of massive aggregation increases to 5 h at 20 °C compared to 2.5 h at 37 °C. Note that the hydrophobic surface treatment used to treat the SPRi and the ATR-FTIR prisms or the coverslip are not exactly the same. A sixteen-carbon moiety was grafted onto gold, but only a six-carbon one was attached onto silicon or glass. Additional SPRi experiments were performed on a gold prism functionalized with an eight-carbon moiety (octyl-thiol) that gave similar results. We therefore believe that the length of the alkyl molecule used to make the material surface hydrophobic does not play a prominent role in HI adsorption and aggregation as long as the contact angle is close to 100°.

Bovine insulin aggregation at acidic pH and high temperature was studied by Bouchard et al. using FTIR [27]. These authors observe a heat-induced conformational transition characterized by the gradual disappearance of the characteristic α-helix peak (1651 cm⁻¹) concomitantly with the appearance of characteristic β-sheet peaks (1670 and 1627 cm⁻¹). In our study, the final HI aggregates obtained upon contact with hydrophobic surfaces present a very similar FTIR signature (Fig. 6E): a major peak at
<figure><img src="image_75.png" /><figcaption>**Fig. 9.** Intermediates in the formation of HI amyloid fibers on hydrophobic surfaces. The scheme summarizes the different steps in the conversion of soluble HI into amyloidal HI aggregates that takes place on hydrophobic surfaces. (1) Soluble insulin adsorption and accumulation on the surface; it is accompanied by changes in the infrared amide A and II' bands. (2) Formation of HI amyloid fibers adsorbed on the surface, characterized by the appearance of the 1708 cm⁻¹ peak, ThT fluorescence and reinforcement of HI binding to the surface. (3) Dissociation of the HI amyloid fibers.</figcaption></figure>

1628 cm⁻¹ (72.1%) and three additional peaks at 1652 cm⁻¹ (16.6%), 1662 cm⁻¹ (2.4%) and 1669 cm⁻¹ (8.9%) (data not shown). The 1628 cm⁻¹ peak fits with the observation that characteristic FTIR β-sheet maxima of protein amyloid fibrils extend from 1611 to 1630 cm⁻¹ [28]. The final insulin aggregates formed at hydrophobic surfaces are therefore similar to the ones formed in solution at high temperature and acidic pH. Nevertheless, Bouchard and co-workers did not characterize any intermediate conformational state, in contrast to our study. We describe a conformation for adsorbed insulin, which has a different FTIR signature than the final HI aggregates (see below).

A clear accumulation of HI on the surface precedes the onset of ThT-positive fluorescence. This accumulation was reported by Ballet et al. [21], but the limited sensitivity and long processing time of the direct biochemical method prevented their documenting it precisely. SPRi experiments show that HI adsorption proceeds in two phases: first, the very fast binding of an HI monolayer, followed by the slow but steady accumulation of HI on top of the monolayer. Assuming that HI adsorbs uniformly, about two protein layers would be present on the surface at the onset of the ThT fluorescence. This signal pattern illustrates the mechanism of insulin aggregation on hydrophobic surfaces: soluble HI is indeed able to bind to adsorbed HI, increasing the local HI concentration tremendously, which is a prerequisite to the formation of aggregates. The precision of SPRi further allows differentiating two HI populations adsorbed on the material surface, characterized by their dissociation kinetics: one strongly bound to the surface, which dissociates slowly, and another one less strongly bound, which dissociates more easily.

Infrared spectroscopy reveals that adsorbed HI is in a conformational state that is distinct from soluble HI and from suspended HI amyloidal aggregates. The amide A and amide II' bands are indeed both shifted to lower energy. It is known that the position of the amide A band, which corresponds to NH stretching, is sensitive to hydrogen bonding [29]. Its shift from 3420 cm⁻¹ to 3276 and 3204 cm⁻¹ thus indicates that adsorbed HI is involved in more hydrogen bonding than soluble HI. The existence of a specific conformation for adsorbed HI is in accordance with Ballet et al. [21], who showed that the bacterial chaperone DnaK recognizes a specific conformation on adsorbed HI. DnaK could bind, though with a diminished affinity, on aggregated HI fibers released in solution, indicating that adsorbed HI and aggregated HI have distinct conformations.

In ATR-FTIR, the amide I band in adsorbed HI is also unique. As shown in Fig. 7, it consists of four peaks, at 1635, 1655, 1687 and 1708 cm⁻¹. Assuming that the assignation of the different amide I peaks provided in the literature for soluble proteins [29,30] also holds for adsorbed proteins, we can interpret the 1635 and 1687 cm⁻¹ peaks as indicative of the presence of β-sheet, and the 1655 cm⁻¹ peak as the residual presence of α-helices. By analogy with the case of cytochrome c [31] and fibroin [32], the 1708 cm⁻¹ peak could be due to intermolecular β-sheets. Furthermore, its shape slowly evolves as HI accumulates over the surface, showing that some maturation takes place (Fig. 7A). Since the kinetics of the 1708 cm⁻¹ peak's appearance roughly matches that of the slow dissociation component evidenced by SPRi, we propose that two states of adsorbed HI are present, corresponding to the fast and slow dissociation components. The first one (HIᵢ), characterized by a prominent peak at 1655 cm⁻¹ (Fig. 7A, HI adsorbed after 1 or 5 min), very similar to that of soluble HI, would dissociate fastest. The second one (HIᵢᵢ), with a characteristic peak at 1708 cm⁻¹ (Fig. 7A, HI adsorbed at 80 min), would dissociate ten times more slowly. Both would be intermediates in the transition from soluble HI to HI amyloid aggregates. Since the HIᵢ → HIᵢᵢ transition is completed in about 30–40 min (Fig. 7), it is tempting to assume that HIᵢᵢ is ThT-positive whereas HIᵢ is not. HIᵢ would thus represent pre-amyloid adsorbed HI and HIᵢᵢ amyloid aggregates (adsorbed HI and amyloid HI in Fig. 9). HIᵢᵢ dissociation from the material surface would allow the formation of mature amyloid fibers in solution. In this state, the infrared signature is different from that of amyloid aggregates bound to the surface, but both are ThT-positive. The mechanical detachment of insulin amyloid fibers has been studied by Ballet et al. [24].

This study underlines the importance of considering the material surface when dealing with protein stability issues. It reveals the complexity of a multi-step adsorption mechanism and the existence of conformationally different protein species that cooperate to form amyloid fibers. Insulin–material interactions are governed by a complex interplay between the hydrophobicity, surface charge and topography of the material. Indeed, it was shown that insulin, although carrying a net negative charge at physiological pH, accumulates in significant amounts on negatively charged, hydrophilic polystyrene sulfonate (water contact angle: 50–60°) [33]. On uncharged poly(ethylene oxide)-coated surfaces (water contact angle: 3.5°), however, no insulin adsorption is measurable [21]. Moreover, Pandey and co-workers have studied the influence of increasing surface hydrophobicity and topography on the nucleation rate using mixed self-assembled monolayers of amine- and octyl-silanes [34]. They show that the lag time of insulin aggregation varies as a function of the amine fraction on the surface and that the morphology of the aggregates depends on the surface chemistry and topography. From these studies we conclude that the conditions that inhibit insulin adsorption on materials and material-induced aggregation, are maximal wettability (water contact angle <5°) and the absence of charges. Further work is clearly needed to examine whether the mechanisms outlined in the case of HI also hold true for other proteins in contact with hydrophobic surfaces, or for HI in contact with other surface chemistries.

## Acknowledgements

This work was supported by the ANR grant "Stabilization of Therapeutical Proteins". L.N. was supported by a PhD grant from the Rhone-Alpes Region (Macodev program). B.J. was an exchange student from Delhi University. P.G. was a summer student from Toronto University, supported by a mobility grant from the Rhone-Alpes Region. We thank Catherine Picart and Flora Gilde for access and help with the ATR-FTIR apparatus. The authors acknowledge the "Pôle de Capteurs Thermométriques et
Calorimétrie" of the Institut Néel laboratory for the use of the oxygen plasma etcher.

## Appendix A. Figures with essential colour discrimination

Certain figures in this article, particularly Figs. 3, 6 and 7, are difficult to interpret in black and white. The full colour images can be found in the on-line version, at http://dx.doi.org/10.1016/j.actbio.2012.09.025.

## References

[1] Westermark P, Benson MD, Buxbaum JN, Cohen AS, Frangione B, Ikeda S, Masters CL, Merlini G, Saraiva MJ, Sipe JD. Amyloid fibril protein nomenclature. J. Amyloid 2002;9:197–200.

[2] Astbury WT, Dickinson S, Bailey K. The X-ray interpretation of denaturation and the structure of the seed globulins. Biochem J 1935;29:2351–2360.1.

[3] Dobson CM. Protein misfolding, evolution and disease. Trends Biochem Sci 1999;24:329–32.

[4] Rudall KM. The proteins of the mammalian epidermis. Adv Protein Chem 1952;7:253–90.

[5] Koo EH, Lansbury PT, Kelly JW. Amyloid diseases: abnormal protein aggregation in neurodegeneration. Proc Natl Acad Sci USA 1999;96:9989–90.

[6] Adams MJ, Blundell TL, Dodson EJ, Dodson GG, Vijayan M, Baker EN, et al. Structure of rhombohedral 2 zinc insulin crystals. Nature 1969;224:491–5.

[7] Feingold V, Jenkins AB, Kraegen EW. Effect of contact material on vibration-induced insulin aggregation. Diabetologia 1984;27:373–8.

[8] Lougheed WD, Albisser AM, Martindale HM, Chow JC, Clement JR. Physical stability of insulin formulations. Diabetes 1983;32:424–32.

[9] Sefton MV, Antonacci GM. Adsorption isotherms of insulin onto various materials. Diabetes 1984;33:674–80.

[10] Lougheed WD, Woulfe-Flanagan H, Clement JR, Albisser AM. Insulin aggregation in artificial delivery systems. Diabetologia 1980;19:1–9.

[11] Dische FE, Wernstedt C, Westermark GT, Westermark P, Pepys MB, Rennie JA, et al. Insulin as an amyloid-fibril protein at sites of repeated insulin injections in a diabetic patient. Diabetologia 1988;31:158–61.

[12] Störkel S, Schneider HM, Müntefering H, Kashiwagi S. Iatrogenic, insulin-dependent, local amyloidosis. Lab Invest 1983;48:108–11.

[13] Moores B, Drolle E, Attwood SJ, Simons J, Leonenko Z. Effect of surfaces on amyloid fibril formation. PLoS One 2011;6:e25954.

[14] Kowalewski T, Holtzman DM. In situ atomic force microscopy study of Alzheimer's beta-amyloid peptide on different substrates: new insights into mechanism of beta-sheet formation. Proc Natl Acad Sci USA 1999;96:3688–93.

[15] Giacomelli CE, Norde W. Influence of hydrophobic Teflon particles on the structure of amyloid beta-peptide. Biomacromolecules 2003;4:1719–26.

[16] McMasters MJ, Hammer RP, McCarley RL. Surface-induced aggregation of beta amyloid peptide by ω-substituted alkanethiol monolayers supported on gold. Langmuir 2005;21:4464–70.

[17] Chi EY, Frey SL, Winans A, Lam KLH, Kjaer K, Majewski J, et al. Amyloid-beta fibrilogenesis seeded by interface-induced peptide misfolding and self-assembly. Biophys J 2010;98:2299–308.

[18] Schladitz C, Vieira EP, Hermel H, Möhwald H. Amyloid-beta-sheet formation at the air-water interface. Biophys J 1999;77:3305–10.

[19] Sluzky V, Tamada J, Klibanov A, Langer R. Kinetics of insulin aggregation in aqueous solutions upon agitation in the presence of hydrophobic surfaces. Proc Natl Acad Sci USA 1991;88:9377–81.

[20] Sluzky V, Klibanov AM, Langer R. Mechanism of insulin aggregation and stabilization in agitated aqueous solutions. Biotechnol Bioeng 1992;40:895–903.

[21] Ballet T, Brukert F, Mangiagalli P, Bureau C, Boulangé L, Nault L, et al. DnaK prevents human insulin amyloid fiber formation on hydrophobic surfaces. Biochemistry 2012;51:2172–80.

[22] Lavalle P, Gergely C, Cuisinier FJG, Decher G, Schaaf P, Voegel JC, et al. Comparison of the structure of polyelectrolyte multilayer films exhibiting a linear and an exponential growth regime: an in situ atomic force microscopy study. Macromolecules 2002;35:4458–65.

[23] Picart C, Ladam G, Senger B, Voegel JC, Schaaf P, Cuisinier FJG, et al. Determination of structural parameters characterizing thin films by optical methods: a comparison between scanning angle reflectometry and optical waveguide lightmode spectroscopy. J Chem Phys 2001;115:1086.

[24] Ballet T, Nault L, Pandey L, Bruckert F, Bureau C, Boulangé L, et al. Insulin adsorbed on hydrophobic surfaces stimulates the formation and release of insulin amyloid fibers. Submitted for publication.

[25] Barth A, Zscherp C. What vibrations tell us about proteins. Q Rev Biophys 2002;35:369–430.

[26] Dzwolak W, Smirnovas V, Jansen R, Winter R. Insulin forms amyloid in a strain-dependent manner: an FT-IR spectroscopic study. Protein Sci 2004;13:1927–32.

[27] Bouchard M, Zurdo J, Nettleton E, Dobson CM, Robinson CV. Formation of insulin amyloid fibrils followed by FTIR simultaneously with CD and electron microscopy. Protein 2000;9:1960–7.

[28] Zandomeneghi G, Krebs MRH, McCammon MG, Fändrich M. FTIR reveals structural differences between native beta-sheet proteins and amyloid fibrils. Protein Sci 2004;13:3314–21.

[29] Kong J, Yu S. Fourier transform infrared spectroscopic analysis of protein secondary structures. Acta Biochim Biophys Sin 2007;39:549–59.

[30] Mizushima S-I, Tsuboi M, Shimanouchi T, Tsuda Y. Spectroscopic investigation of the strength of hydrogen bonds formed by amides. Spectrochim Acta 1955;7:100–7.

[31] Marboutin L, Boussac A, Berthomieu C. Redox infrared markers of the heme and axial ligands in microperoxidase: bases for the analysis of c-type cytochromes. J Biol Inorg Chem 2006;11:811–23.

[32] Hu X, Kaplan D, Cebe P. Determining beta-sheet crystallinity in fibrous proteins by thermal analysis and infrared spectroscopy. Macromolecules 2006;39:6161–70.

[33] Jeworrek C, Hollmann O, Steitz R, Winter R, Czeslik C. Interaction of IAPP and insulin with model interfaces studied using neutron reflectometry. Biophys J 2009;96:1115–23.

[34] Pandey LM, Le Denmat S, Delabouglise D, Bruckert F, Pattanayek SK, Weidenhaupt M. Surface chemistry at the nanometer scale influences insulin aggregation. Colloids Surf B 2012;100C:69–76.
# Peptides that form β-sheets on hydrophobic surfaces accelerate surface-induced insulin amyloidal aggregation

<figure><img src="image_76.png" /></figure>

CrossMark

Laurent Nault<sup>a</sup>, Charlotte Vendrely<sup>a</sup>, Yves Bréchet<sup>b</sup>, Franz Bruckert<sup>a</sup>, Marianne Weidenhaupt<sup>a,*</sup>

## ARTICLE INFO

**Article history:**

Received 12 July 2012

Revised 19 October 2012

Accepted 16 November 2012

Available online 16 March 2013

Edited by Jesus Avila

**Keywords:**

Material surface

Protein aggregation

Amyloidogenic peptide

Insulin

## ABSTRACT

Interactions between proteins and material or cellular surfaces are able to trigger protein aggregation in vitro and in vivo. The human insulin peptide segment LVEALYL is able to accelerate insulin aggregation in the presence of hydrophobic surfaces. We show that this peptide needs to be previously adsorbed on a hydrophobic surface to induce insulin aggregation. Moreover, the study of different mutant peptides proves that its sequence is less important than the secondary structure of the adsorbed peptide on the surface. Indeed, these pro-aggregative peptides act by providing stable β-sheets to incoming insulin molecules, thereby accelerating insulin adsorption locally and facilitating the conformational changes required for insulin aggregation. Conversely, a peptide known to form α-helices on hydrophobic surfaces delays insulin aggregation.

**Structured summary of protein interactions:**

HI and HI bind by fluorescence technology (View interaction)

© 2013 Federation of European Biochemical Societies. Published by Elsevier B.V. All rights reserved.

## 1. Introduction

Many protein solutions are prone to aggregation in vitro. This is particularly critical for therapeutic proteins because aggregate formation decreases drug activity and can induce immunogenic reactions when injected in patients. For instance, the most used protein hormone, insulin, can be found in fibrillar form at the site of frequent insulin injections, which causes injection amyloidosis [1]. Moreover, the increasing prevalence of human diseases, characterized by the presence of large amounts of aggregated proteins, like amyloidosis, Alzheimer's, Parkinson's and prion diseases, requires mechanistic studies to understand protein aggregate formation.

The mechanism of protein fibrillation is generally modeled in three steps. (1) Conformational changes, due to defects in the cellular folding processes or due to an abnormal protein environment. (2) Nucleation, during which the misfolded individual proteins are stabilized, mostly by intermolecular interactions, which drives the formation of oligomers. Such oligomers can then evolve into stable nuclei. (3) Growth, during which the nuclei will grow upon binding of other misfolded proteins, leading to the formation of large fibrils. In vivo, the growing step has a major impact on cell survival [2].

Human insulin (HI) has often been used as a model protein in fibrillation studies as it is a non-expensive protein which fibrillates

rapidly in vitro, particularly at low pH, high temperature, high ionic strength and on hydrophobic surfaces [3–5]. In a preceding study [6], we have shown that the presence of hydrophobic surfaces is essential for HI fibrillation at pH 7.3 and 37 °C. Such surfaces were assumed to induce the unfolding of HI monomers adsorbed on them, but it appears that all three steps, including nucleation and fibril elongation also take place on the surface, leading eventually to the release of mature fibrils into solution.

Different studies show that the onset of aggregate nucleation can be accelerated by the addition of peptides, which present a pro-aggregative activity [7–10]. Such amyloidogenic peptides are mostly hydrophobic, have beta-sheet conformation and are capable to fibrillate by themselves. They are commonly derived from protein sequences suspected to be involved in fibrillation. In 2009, Ivanova et al. [11] showed that a 7 amino-acid peptide sequence, present in the B chain of native HI (LVEALYL, residues B11–B17), was able, at sub-stoichiometric concentrations, to accelerate the nucleation step of human HI at pH 2.5. On the other hand, it is known that surface hydrophobicity has an important effect on the HI nucleation rate [12] and, given the hydrophobic nature of this peptide, its amyloidogenic properties are likely to rely on its interaction with hydrophobic surfaces. We recently demonstrated that the LVEALYL peptide-driven acceleration of HI aggregation is strictly dependent on the presence of hydrophobic surfaces [6].

In this study, we use mutants of the LVEALYL peptide together with HI to investigate the effects of amino acid changes in this

<footnote><sup>a</sup>Laboratoire des Matériaux et du Génie Physique, Phelma-Minatec, 3 Parvis Louis Néel CS 50257, F-38016 Grenoble Cedex 1, France</footnote>

<footnote><sup>b</sup>Laboratoire de Science et Ingénierie des Matériaux et des Procédés, Phelma-Campus, 1130 rue de la Piscine, BP 75, F-38402 Saint-Martin d'Hères Cedex, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: marianne.weidenhaupt@grenoble-inp.fr (M. Weidenhaupt).</footnote>
sequence on HI aggregation kinetics, protein adsorption on hydrophobic surfaces and changes in peptide secondary structure. When compared to previous studies measuring the effects of amyloido- genic peptides on HI aggregation kinetics, we have introduced two novel and important parameters: first, the experiments are done at physiological pH and, second, the effect of material surface hydrophobicity is analyzed. Our work suggests that the LVEALYL peptide and mutants thereof induce HI aggregation only if they are previously adsorbed on the material surface. Moreover, it appears that their primary sequence is less important than their ability to stably adsorb in beta-structure on the surface.

## 2. Materials and methods

### 2.1. Chemicals

If not otherwise stated, all chemicals were purchased from Sigma-Aldrich. Experiments were conducted in TBS (25 mM Tris-HCl, pH 7.3, 125 mM NaCl and 2 mM MgCl₂). HI (recombinant human HI, expressed in yeast) solutions were prepared at 0.5 mg mL⁻¹ (86 µM). All solutions were filtered (0.22 µm) before use. Peptides were purchased from Genecust (Luxembourg) and a concentrated stock solution was made at 860 µM in 20 mM NaOH.

### 2.2. Monitoring of aggregation kinetics

HI aggregation assays were conducted as 8 replicates in plastic 96-well microplates. Polystyrene (Greiner Bio-One, water contact angle = 85° ± 4.7), or PEO-coated, (Corning 3651 microplate, water contact angle = 3.5° ± 5.8) microplates were used. In fluorescence assays, black polystyrene microplates were used (Nunc Nunclon® Δ Surface). The plates were covered by plastic sheets, incubated at 37 °C and shaken at 1200 rpm (Heidolph Titramax, 1.5 mm vibration orbit). Thioflavin T (ThT, 20 µM) fluorescence was directly measured (λ<sub>ex</sub> = 450 nm, λ<sub>em</sub> = 482 nm) in the wells [6].

### 2.3. Kinetic analysis

The aggregation kinetics proceeds in three phases: a lag phase, where the signal is not statistically different from the baseline (mean ± standard deviation), a linear growth phase and a plateau phase. Experimentally, the lag time was defined by the intercept between the linear growth phase and the baseline. The growth rate was defined as the slope of the linear phase and the plateau as the maximum value attained. The parameters were calculated on individual kinetics corresponding to different replicates, and the given statistics represent the average and the standard deviation for each parameter.

### 2.4. Surface seeding using peptides

96-Well microplates were filled with peptides diluted at 8.6 µM in TBS and incubated at 37 °C, 1200 rpm for 10 min. Wells were then washed once with TBS and filled with HI (86 µM) and incubated at 37 °C, 1200 rpm. HI aggregation kinetics was monitored using ThT as described.

### 2.5. Quantification of adsorbed protein

HI (86 µM) and/or peptide solutions (8.6 µM) were incubated in 96-well hydrophobic microplates in TBS for 30 min at 37 °C, 1200 rpm. Wells were then washed with TBS and adsorbed peptides and protein were desorbed using Sodium Dodecyl Sulfate (5%) for 30 min at 37 °C and 1200 rpm. The peptide and/or protein concentrations were determined using the Bicinchoninic Assay using Bovine Serum Albumin as a standard.

### 2.6. Adsorption kinetics and infrared spectroscopy

Peptide and HI association and dissociation kinetics were studied on PEG- and C₁₆-coated surfaces by SPRi as described in Nault et al. [13]. The peptide conformation in solution and adsorbed on hydrophobic surfaces was determined by FTIR transmission or ATR-FTIR experiments, respectively [13].

## 2.7. Estimation of the hydrophobic contribution to the peptide binding energy to hydrophobic surfaces

The hydrophobic contribution to the binding energy of 7 amino acid long peptides with a hydrophobic surface was calculated for two β-sheet configurations by the following formula:

$$ \Delta G_{\text{conf1}} = \sum_{1.3,5,7} \Delta G_i - \sum_{2.4,6} \Delta G_i \quad (1) $$

$$ \Delta G_{\text{conf2}} = \sum_{2.4,6} \Delta G_i - \sum_{1.3,5,7} \Delta G_i \quad (2) $$

$$ \Delta G_{\text{sol}} = \sum_{1}^{7} \Delta G_{i} \quad (3) $$

where ΔGᵢ is the free transfer energy of the ith amino acid side chain [14]. Conformation 1 and 2 correspond to the exposure of even or odd amino acids to solvent, respectively. The third calculation corresponds to the hydrophobic contribution of the peptide transfer energy into solution, with all amino acids exposed to the solvent. This calculation does not consider the contribution of N- and C-termini, which is identical for all peptides and conformations. In the interpretation of the data, peptides are assumed to adopt the conformation having the lowest interaction energy.

## 3. Results

### 3.1. The LVEALYL peptide accelerates HI aggregation kinetics when bound to hydrophobic surfaces

As shown before [6], the LVEALYL peptide accelerates HI aggregation kinetics at pH 7.3 and 37 °C, only in the presence of hydrophobic surfaces (polystyrene) but not in the presence of PEG-coated surfaces preventing HI binding. The maximum effect is observed at a peptide concentration of 8.6 µM, and the lag time is reduced by 66% on average (Table 1). We quantified the peptide adsorption on microwells. Table 1 shows that 0.06 µg LVEALYL peptide remains adsorbed on the hydrophobic surface after 30 min, corresponding to 4.5% of the initial peptide amount. These minute amounts have a dramatic effect on HI aggregation, as shown by the following seeding experiment. Hydrophobic surfaces were pre-incubated for 10 min with peptide, washed and exposed to HI. Peptide preadsorption decreases the lag-time of HI aggregation by 63% (Table 1). It is noteworthy that preincubating the surface with HI does not decrease the lag time (Table 1, line1). Moreover, supplying the solution with 0.06 µg LVEALYL peptide, does not affect HI aggregation kinetics. The pro-aggregative effect of the peptide is therefore due to the adsorbed fraction on the surface. Upon adsorption, but not in solution, the LVEALYL peptide presents molecular features that strongly accelerate HI binding and aggregation.

### 3.2. The pro-aggregative peptides LVEALYL and LVAELYL have a strong effect on HI aggregation and adsorption kinetics

The strong effect of the LVEALYL peptide on HI aggregation kinetics prompted us to study different peptide variants (Table 1). All experiments were done at the same peptide (8.6 µM) and HI (86 µM) concentrations. None of the peptides studied here
<caption>**Table 1**</caption>

Physico-chemical and pro-aggregative properties of different peptides. The binding energy corresponds to the hydrophobic contribution to the minimum interaction energy of the peptide bound to the surface, calculated by Eqs. (1) or (2) (column 2). For the (LKKLLKL)$_2$ peptide no interaction energy was calculated since this peptide is in α-helical configuration. Lag times were determined from the aggregation kinetics of a 1:10 peptide:protein mixture (column 3) and for a surface first exposed to the indicated peptide (8.6 µM) for 10 min, washed and then incubated with a fresh HI solution (86 µM) (column 4). Adsorption of peptides and peptide plus HI mixtures on hydrophobic surfaces (2 cm²) was determined as described in Section 2 (column 5 and 6).

<table><thead><tr><th rowspan="2">Peptide sequence</th><th rowspan="2">Interaction energy (kcal mol<sup>-1</sup>)</th><th colspan="3">Lag time of HI aggregation kinetics (h)</th><th colspan="2">Protein adsorbed on the surface after 30 min (µg)</th></tr><tr><th>8.6 µM peptide mixed with 86 µM HI</th><th>After 10 min seeding</th><th>8.6 µM peptide alone</th><th>8.6 µM peptide mixed with 86 µM HI</th></tr></thead><tbody><tr><td>No peptide</td><td></td><td>4.4 ± 0.4</td><td>4.3 ± 0.4<sup>a</sup></td><td></td><td>0.19 ± 0.01</td><td></td></tr><tr><td>LVEALYL</td><td>-3.3</td><td>1.5 ± 0.2</td><td>1.7 ± 0.3</td><td>0.06 ± 0.01</td><td>0.21 ± 0.01</td><td></td></tr><tr><td>LVAELYL</td><td>-16.3</td><td>0.4 ± 0.1</td><td>2.3 ± 0.3</td><td>0.10 ± 0.01</td><td>4.28 ± 0.10</td><td></td></tr><tr><td>LSASLSL</td><td>-8.2</td><td>0.5 ± 0.1</td><td>2.5 ± 0.3</td><td>0.13 ± 0.01</td><td>0.25 ± 0.01</td><td></td></tr><tr><td>SVSASYS</td><td>-1.1</td><td>2.0 ± 0.1</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr><tr><td>LPEALYL</td><td>-0.5</td><td>4.1 ± 0.3</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr><tr><td>LVEPLYL</td><td>-1.5</td><td>4.2 ± 0.2</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr><tr><td>SVSPSYS</td><td>-0.7</td><td>4.3 ± 0.4</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr><tr><td>LVTALYL</td><td>-6.1</td><td>0.5 ± 0.1</td><td>2.7 ± 0.3</td><td>0.16 ± 0.04</td><td>1.6 ± 0.2</td><td></td></tr><tr><td>LVATLYL</td><td>-6.9</td><td>0.6 ± 0.1</td><td>1.8 ± 0.3</td><td>0.15 ± 0.02</td><td>0.5 ± 0.2</td><td></td></tr><tr><td>LVTPLYL</td><td>-7.9</td><td>4.3 ± 0.1</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr><tr><td>LVPTLYL</td><td>-5.1</td><td>4.6 ± 0.1</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr><tr><td>ISISISI</td><td>-10.6</td><td>0.4 ± 0.1</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr><tr><td>(LKKLLKL)$_2$</td><td>ND</td><td>>30</td><td>ND</td><td>ND</td><td>ND</td><td></td></tr></tbody></table>

aggregates on its own, nor triggers HI aggregation in the presence of PEG-coated surfaces, confirming that the kinetic enhancement is dependent on the presence of hydrophobic surfaces. Among the peptides studied, the swapped peptide LVAELYL, where the third and fourth amino acid are exchanged, exhibits a remarkably short lag time (0.4 h). This peptide binds also strongly to hydrophobic surfaces (0.1 µg) and pre-adsorption reduces the lag time of HI aggregation down to 2.3 h (Table 1).

The binding kinetics of both peptides, alone and in combination with HI, were studied by SPRI (Fig. 1A and B). As shown before [13], HI alone binds specifically to hydrophobic surfaces, in a biphasic manner. Fast, reversible binding is followed by a slower increase of HI adsorption, resulting in the formation of a strongly bound HI pool ([13] and Fig. 1). The LVEALYL and LVAELYL peptides alone bind weakly to the hydrophobic surface (Fig. 1A and B). Mixing one of the peptides with HI enhances protein adsorption on the hydrophobic surface but has no effect on HI adsorption on the hydrophilic surface (Fig. 1A and B). More specifically, upon injection of HI with either peptide, there is an increase of the fast association component (first minutes of Fig. 1A and B) and of the slow dissociation component of the SPRI signal (last 10 min of Fig. 1A and last 20 min of Fig. 1B), compared to HI alone. The addition of sub-stoichiometric amounts of both peptides to HI therefore induces the formation of strongly adsorbed HI on the hydrophobic surface. A direct measurement of protein binding to microwell plastic surfaces confirms this result (Table 1). In the presence of the swapped peptide, the total protein bound is larger than in the presence of HI or peptide alone. The wild-type peptide also enhances, but to a lesser extent, protein binding on the surface (Table 1).

### 3.3. The wild type and the swapped peptides adopt a β-sheet conformation on hydrophobic surfaces

Infrared spectroscopy is sensitive to protein secondary structure in the amide I band and was used to analyze the conformational state of different peptides in solution (FTIR) or adsorbed on hydrophobic surfaces (ATR-FTIR). Decomposing the band into peaks shows that in solution, the wild type and swapped peptides adopt mostly unstructured conformations (Fig. 2A and B, left panels). Indeed, they present a major peak around 1645 cm⁻¹, indicative of random coil conformation, which accounts for 37% and 32% in the wildtype and swapped peptides' solution spectra, respectively (Table 1S). After 30 min adsorption on hydrophobic surfaces,

<figure><img src="image_77.png" /></figure>

<figure><img src="image_78.png" /><figcaption>**Fig. 1.** SPRI sensorgrams of protein adsorption on hydrophobic surfaces. The protein solutions (8.6 µM peptide, 86 µM HI or a mixture of 8.6 µM peptide and 86 µM HI) were injected at *t* = 0 min and the association kinetics were monitored for 10 min (A) or 15 min (B). The surface was then washed with buffer and the protein dissociation kinetics were monitored for 15 min (A) or 20 min (B). The signal represented is the difference between the signal recorded on the C₁₆-coated side and the PEG-coated side of the prism. (A) SPRI curves obtained with peptides LVEALYL and LSASLSL, (B) SPRI curves obtained with peptide LVAELYL.</figcaption></figure>

the amide I band of both peptides changes and reveals prominent peaks around 1694, 1680, 1664, 1648, 1633 and 1622 cm⁻¹ (Fig. 2A and B, right panels). This indicates that the conformation of the adsorbed wild type and swapped peptides consists predominantly of

<footnote>a Seeding with 86 µM HI.</footnote>
internal and intermolecular β-sheets and β-turns, the residual unstructured coil contribution consisting of 17 and 22%, respectively (Table 1S).

### 3.4. Characterization of other amyloidogenic peptides: influence of primary and secondary structure

In order to explain the peptide effect on the HI aggregation rate, a simple model was built, assuming that amyloidogenic peptides adopt a β-sheet conformation parallel to the material surface, stabilized by intermolecular hydrogen bonds and side chain interactions with the hydrophobic surface on one side, and the solvent on the other side. Amino acid side chains indeed alternatively point to one side or the other of a β-strand. For a 7 amino acid long peptide, two such conformations exist, with odd or even amino acids exposed to solvent. For each conformation, the hydrophobic contribution of amino acid side chains to the binding energy was summed up using the transfer energies from benzene to water, amino acids exposed to solvent contributing positively, while those in contact with the surface contributing negatively Eqs. (1) and (2). In this way, one can determine the energetically most favorable conformation of a given peptide adsorbed on a hydrophobic surface. For a stable adsorption, this energy should be lower than the one for the peptide in solution Eq. (3). For the LVEALYL peptide, the most stable β-sheet conformation exposes the L-E-L-L amino acids towards the solution, and the -V-A-Y- amino acids towards the surface (interaction energy = -3.3 kcal mol⁻¹). For the LVAEYL peptide, the most stable conformation exposes the -V-E-Y- amino acids towards the solution, and the L-A-L-L amino acids towards the surface (interaction energy = -16.3 kcal mol⁻¹). This justifies the better stability of the swapped peptide.

Two other peptides were studied, SVSASYS and LSASLSL, based on the wild-type and the swapped peptides respectively, where the amino acids supposed to face the surface were conserved and the ones turned to the solvent replaced by serines (S). For both peptides, the most stable conformation exposes hydrophilic S to the solution and hydrophobic amino acids to the surface (interaction energy = -8.2 kcal mol⁻¹ for LSASLSL and -1.1 kcal mol⁻¹, for SVSAVYS). In the HI aggregation assay, the SVSASYS and the LSASLSL peptides behaved similarly to their parent peptides (Table 1), showing that the primary sequence exposed to the solvent was not critical. The stronger effect of the LSASLSL peptide, compared to the SVSASYS one, is again justified by its higher stability on hydrophobic surfaces. The LSASLSL peptide also presents a β-sheet rich conformation when adsorbed on hydrophobic surfaces (Fig. 2C and Table 1S), and enhances HI binding on the surface (Fig. 1).

To confirm the importance of peptide conformation on the surface, the (LKKLLKL)₂ peptide, known to form α-helices on hydrophobic surfaces [15], was tested. This peptide indeed proved to have completely different FTIR spectra, when compared to the wildtype or swapped peptides (Fig. 2D, left and right panels). Its transmission spectrum indicates that the (LKKLLKL)₂ peptide adopts β-sheet, random coil and α-helix conformations in solution (60%, 7% and 33%, respectively; Table 1S). Strikingly, when adsorbed on a hydrophobic surface, its ATR-FTIR signature reveals typical peaks around 1641 and 1655 cm⁻¹, which indicates that this peptide is predominantly in α-helical and random conformation (Table 1S) upon adsorption. At substoichiometric concentrations, (LKKLLKL)₂ delays HI aggregation for up to 30 h (Table 1). We therefore conclude that the conformation of the adsorbed (LKKLLKL)₂ peptide is not pro-aggregative but rather stabilizes HI.

Furthermore a proline residue (P), was introduced to disrupt the wildtype peptide secondary structure: LPEALYL, LVEPLYL and SVSPSYS. None of these three peptides affect HI aggregation

kinetics (Table 1), which consolidates the importance of peptide β-sheet conformation on the surface.

Replacing the negatively charged glutamate residue (E) by a threonine (T) in the wild type or swapped peptide had no effect on the pro-aggregative property of the peptides, showing that electrostatic interactions were not determinant (LVTALYL and LVATLYL in Table 1). Again, the additional A to P mutation abolished the pro-aggregative effect of these peptides (LVTPLYL and LVPTLYL in Table 1).

Finally, we analyzed a peptide, ISISISI, exposing a series of three Ser (S) to the solution and being adsorbed via four Ile (I) to the hydrophobic surface. This peptide has a sequence that is similar to the LKLKLKL peptide, known to form β-sheets on hydrophobic surfaces [15–17], and has no common residues with LVEALYL. The HI aggregation lagtime, obtained using this peptide in seeding experiments, is 0.4 h. This peptide therefore also shows a pro-aggregative effect on HI-surface aggregation and confirms that the primary sequence is not paramount.

### 4. Discussion

In this study, we show that the LVEALYL peptide, adsorbed on hydrophobic surfaces at sub-stoichiometric concentrations, favors HI binding and the formation of HI amyloidal fibers. We previously showed that HI adsorbed on hydrophobic surfaces exposes a mobile containing the LVEALYL peptide, which is recognized by the DnaK/DnaJ bacterial chaperones [6]. Combining these two observations indicates how HI amyloidal aggregates form on hydrophobic surfaces. HI molecules adsorb and change their conformation, exposing the LVEALYL peptide that allows further HI binding in a β-sheet conformation. The conformational change induced on HI directly adsorbed on the surface thus propagates to incoming HI that binds to adsorbed HI. This mechanism is in accordance with the insulin amyloid fiber model proposed by Ivanova et al. [11]. There is some similarity with the prion self-replication mechanism, with soluble and adsorbed HI playing the role of PrP<sup>C</sup> and PrP<sup>SC</sup>, respectively. An important difference is that HI amyloid aggregates released into solution do not seed efficiently fiber growth.

This work confirms the central role played by the LVEALYL peptide, pointed out by Ivanova et al. [11]. We confirm that the SLYQL-ENY peptide, another HI segment putatively involved in the HI amyloid structure, does not stimulate HI aggregation in contact with hydrophobic material surfaces. This is reasonable since this peptide is globally hydrophilic ($ΔG_{sol} = -12.3$ kcal mol⁻¹) and therefore not likely to adopt a β-sheet conformation on hydrophobic surfaces. Interestingly, introducing two mutations so that hydrophobic amino acids are placed at even locations (LYQLEAY peptide, $ΔG_{conf1} = -13.9$ kcal mol⁻¹) has a mild but significant effect on HI aggregation kinetics (lag time = 3.4 h ± 0.1). Our study, although limited to a subset of all possible peptides, strongly suggests that the interaction of HI with the adsorbed peptide is not highly sensitive to the primary structure, as long as the adsorbed peptide provides a stable β-structure on the surface. Furthermore, the effect of the short peptides on insulin aggregation is related to their surface binding energy (minimum -5 kcal mol⁻¹) and requires a β-sheet conformation (Fig. 3). The interaction with the peptide-borne β-structure drives stable HI adsorption by triggering an α to β transition in HI and consequent β-sheet formation between neighboring peptides and incoming HI molecules. This effect is reminiscent of the effect of silver iodide crystals on super-cooled H₂O that induces the formation of ice crystals because of a match between the crystal structures of β-Agl and ice. It is already known that short peptides are able to influence full-length protein amyloid aggregation at iso-stoichiometric concentrations [7–9]. Nevertheless, the impact of container surface chemistry is not systematically taken into account, despite the fact that hydrophobic
<figure><img src="image_79.png" /><figcaption>**Fig. 2.** Amide I band of the LVEALYL (A), LVAELYL (B) and LSASLSL (C) and (LKKLLKL)₂ peptides in solution (left) and adsorbed on hydrophobic surfaces (right). Transmission FTIR spectra were recorded with 8.6 µM peptides in deuterated buffer. For ATR-FTIR, peptides were introduced at 8.6 µM in the flow chamber of a hexylmethyl-coated silicon prism. After 30 min, the prism surface was washed with buffer and the infrared spectra were recorded. Thick line: amide I band after linear baseline correction. Thin lines: decomposition of the amide I band into individual peaks.</figcaption></figure>
<figure><img src="image_80.png" /><figcaption>**Fig. 3.** Effect of different peptides on insulin kinetics. Aggregation kinetics of HI (86 µM) alone or mixed with 8.6 µM peptide were determined. The nucleation rates *k* (in the presence of peptide) and *k*₀ (in the absence of peptide) were calculated as the inverse of the lag time. For each peptide, the *k*/*k*₀ ratio is plotted as a function of the binding energy of the peptide on hydrophobic surfaces in β-strand conformation Eqs. (1) or (2). Circles: peptides without proline residue. Squares: peptides containing one proline residue. Data are from Table 1. Lines are hand drawn.</figcaption></figure>

inter interfaces are known to be able to induce the aggregation of differ- ent proteins [5,18]. Here we show that peptides present in sub- stoichiometric concentrations, adsorbed on hydrophobic surfaces are able to accelerate or delay HI aggregation.

Acknowledgements

This work was supported by the ANR Grant "Stabilization of Therapeutical Proteins". Laurent Nault held a doctoral fellowship from la Région Rhône-Alpes. We thank Catherine Picart and Flora Gilde for access to and assistance with the ATR-FTIR. The authors acknowledge the "Pôle de Capteurs Thermométriques et Calorimétrie" of the Institut Néel for the use of the oxygen plasma etching equipment.

**Appendix A. Supplementary data**

Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.febslet.2012. 11.036.

**References**

[1] Dische, F.E., Wernstedt, C., Westermark, G.T., Westermark, P., Pepys, M.B., Rennie, J.A., Gilbey, S.G. and Watkins, P.J. (1988) Insulin as an amyloid-fibril protein at sites of repeated insulin injections in a diabetic patient. Diabetologia 31, 158–161.

[2] Wogulis, M., Wright, S., Cunningham, D., Chilcote, T., Powell, K. and Rydel, R.E. (2005) Nucleation-dependent polymerization is an essential component of amyloid-mediated neuronal cell death. J. Neurosci. 25, 1071–1080.

[3] Brange, J., Andersen, L., Laursen, E.D., Meyn, G. and Rasmussen, E. (1997) Toward understanding insulin fibrillation. J. Pharm. Sci. 86, 517–525.

[4] Ahmad, A., Uversky, V.N., Hong, D. and Fink, A.L. (2005) Early events in the fibrillation of monomeric insulin. J. Biol. Chem. 280, 42669–42675.

[5] Sluzky, V., Tamada, J., Klibanov, A. and Langer, R. (1991) Kinetics of insulin aggregation in aqueous solutions upon agitation in the presence of hydrophobic surfaces. Proc. Natl. Acad. Sci. USA 88, 9377–9381.

[6] Ballet, T., Brukert, F., Mangiagalli, P., Bureau, C., Boulangé, L., Nault, L., Perret, T. and Weidenhaupt, M. (2012) DnaK prevents human insulin amyloid fiber formation on hydrophobic surfaces. Biochemistry 51, 2172–2180.

[7] Scrocchi, L.A., Ha, K., Chen, Y., Wu, L., Wang, F. and Fraser, P.E. (2003) Identification of minimal peptide sequences in the (8–20) domain of human islet amyloid polypeptide involved in fibrillogenesis. J. Struct. Biol. 141, 218–227.

[8] Du, H.-N., Li, H.-T., Zhang, F., Lin, X.-J., Shi, J.-H., Shi, Y.-H., Ji, L.-N., Hu, J., Lin, D.-H. and Hu, H.-Y. (2006) Acceleration of alpha-synuclein aggregation by homologous peptides. FEBS Lett. 580, 3657–3664.

[9] Kim, J.R. and Murphy, R.M. (2004) Mechanism of accelerated assembly of beta-amyloid filaments into fibrils by KLVFFK(6). Biophys. J. 86, 3194–3203.

[10] Hong, D.-P. and Fink, A.L. (2005) Independent heterologous fibrillation of insulin and its B-chain peptide. Biochemistry 44, 16701–16709.

[11] Ivanova, M.I., Sievers, S.A., Sawaya, M.R., Wall, J.S. and Eisenberg, D. (2009) Molecular basis for insulin fibril assembly. Proc. Natl. Acad. Sci. USA 106, 18990–18995.

[12] Sluzky, V., Klibanov, A.M. and Langer, R. (1992) Mechanism of insulin aggregation and stabilization in agitated aqueous solutions. Biotechnol. Bioeng. 40, 895–903.

[13] Nault, L., Guo, P., Jain, B., Bréchet, Y., Bruckert, F. and Weidenhaupt, M. (2013) Human insulin adsorption kinetics, conformational changes and amyloidal aggregate formation on hydrophobic surfaces. Acta Biomater. 9, 5070–5079.

[14] Engelman, D.M., Steitz, T.A. and Goldman, A. (1986) Identifying nonpolar transbilayer helices in amino acid sequences of membrane proteins. Annu. Rev. Biophys. Biophys. Chem. 15, 321–353.

[15] DeGrado, W.F. and Lear, J.D. (1985) Induction of peptide conformation at apolar water interfaces. 1. A study with model peptides of defined hydrophobic periodicity. J. Am. Chem. Soc. 107, 7684–7689.

[16] Collier, G., Vellore, N.A., Yancey, J.A., Stuart, S.J. and Latour, R.A. (2012) Comparison between empirical protein force fields for the simulation of the adsorption behavior of structured LK peptides on functionalized surfaces. Biointerphases 7, 24.

[17] Weidner, T., Apte, J.S., Gamble, L.J. and Castner, D.G. (2010) Probing the orientation and conformation of alpha-helix and beta-strand model peptides on self-assembled monolayers using sum frequency generation and NEXAFS spectroscopy. Langmuir 26, 3433–3440.

[18] McMasters, M.J., Hammer, R.P. and McCarley, R.L. (2005) Surface-induced aggregation of beta amyloid peptide by ω-substituted alkanethiol monolayers supported on gold. Langmuir 21, 4464–4470.
# Dual Effect of (LK)nL Peptides on the Onset of Insulin Amyloid Fiber Formation at Hydrophobic Surfaces

Karim Chouchane, Charlotte Vendrely,† Myriam Amari, Katie Moreaux, Franz Bruckert, and Marianne Weidenhaupt*

**ABSTRACT:** Soluble proteins are constantly in contact with material or cellular surfaces, which can trigger their aggregation and therefore have a serious impact on the development of stable therapeutic proteins. In contact with hydrophobic material surfaces, human insulin aggregates readily into amyloid fibers. The kinetics of this aggregation can be accelerated by small peptides, forming stable beta-sheets on hydrophobic surfaces. Using a series of (LK)nL peptides with varying length, we show that these peptides, at low, substoichiometric concentrations, have a positive, cooperative effect on insulin aggregation. This effect is based on a cooperative adsorption of (LK)nL peptides at hydrophobic surfaces, where they form complexes that help the formation of aggregation nuclei. At higher concentrations, they interfere with the formation of an aggregative nucleus. These effects are strictly dependent on the their adsorption on hydrophobic material surfaces and highlight the importance of the impact of materials on protein stability. (LK)nL peptides prove to be valuable tools to investigate the mechanism of HI aggregation nuclei formation on hydrophobic surfaces.

<figure><img src="image_81.png" /></figure>

## ■ INTRODUCTION

Protein stability is a crucial parameter, foremost because it is a prerequisite for cellular function but also for safety and economic reasons in the production, storage, and delivery of therapeutics. Alteration of the protein structure usually entails the formation of aggregates, among which the amyloid fiber is widely studied. Amyloid aggregation of proteins in vivo is linked to major diseases, like Alzheimer's or Parkinson's.¹ Insulin, the most produced therapeutic protein, is known for decades for its propensity to aggregate into fibers, which is a serious drawback for the development of novel diabetes therapies.² Understanding the mechanisms involved in protein fibrillation is therefore essential to be able to control protein stability.

The transition from the soluble protein to the aggregated amyloid fiber is generally described in three successive steps: (1) a conformational change triggered by improper folding or environmental conditions, (2) stabilization of partly unfolded protein monomers by intermolecular interactions to form nucleation sites, and (3) association of successive unfolded monomers with the nuclei to promote fiber growth. Accordingly, a lag phase (steps 1 and 2) and a growth phase (step 3) are characteristically defined to describe protein fibrillation. For human insulin (HI), fibrillation can be triggered in vitro by low pH, high temperature, high ionic strength, and hydrophobic surfaces.³⁻⁵ We have shown that at pH 7 and 37 °C, all three steps of the fibrillation process are strictly dependent on the presence of hydrophobic surfaces.⁶ This finding highlights an aspect that is generally overseen in protein aggregation studies: the impact of the container material used

for the experiments. In most cases, plastic containers are used and their hydrophobic surface represents both the site and trigger for insulin aggregation. In the following, we define nuclei as the surface-bound structures, able to trigger fibril growth (seeds). All complexes present during the maturation of these nuclei are termed prenuclei.

In an attempt to understand protein aggregation into amyloid fibers, multiple studies have been aimed at identifying minimal peptide sequences, inhibiting, or accelerating the aggregation kinetics of the parent protein. For example, an 8 amino-acid prion peptide segment (residues 113–120: AGAAAAGA), a 6 amino-acid modified peptide from Aβ (residues 16–20: KLVFF) and a 5 amino-acid segment from IAPP (residues 22–27: NFGAIL) can delay fibril formation of their respective full-length protein.⁷⁻¹⁰ Such peptides are often hydrophobic, adopt a beta-sheet conformation, and fibrillate by themselves. The inhibitory effect is generally obtained at high concentrations, ranging from equimolar to concentrations exceeding ten times the protein concentration. On the other hand, amyloidogenic peptides, accelerating the aggregation lag phase, were also discovered.¹¹⁻¹³ They are often effective at low concentrations compared to the protein. Interestingly, many peptides have both accelerating and inhibiting effects on protein aggregation, depending on their relative concentrations. Ivanova et al. have shown that the HI B-chain peptide LVEALYL (residues B11 to B17) is able, at substoichiometric

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<footnote>†University Grenoble Alpes, CNRS, LMGP, F-38000 Grenoble, France</footnote>

<footnote>‡Cergy Pontoise University, ERRMECe, I-MAT FD4122, F-95302 Cergy Pontoise, France</footnote>
concentrations to reduce the HI aggregation lag-time significantly and, at higher concentrations, to inhibit fibril growth.¹²

Using the same LVEALYL peptide and mutants thereof, we could demonstrate that the amyloidogenic effect described by Ivanova et al.¹² depends on the adsorption of the peptide in beta-sheet conformation on a hydrophobic surface.¹⁴ Moreover, we could show that the secondary structure and not the sequence of the adsorbed peptide was crucial.¹⁴,¹⁵

Amyloidogenic peptides thus represent a tool to elucidate how a hydrophobic surface can trigger the formation of aggregation nuclei and hence amyloid fiber growth. They can be customized in sequence and chemical properties (functional groups, fluorescent labels). In this way, they are at the same time amyloid aggregation inducers and tools to investigate the aggregates they trigger. In order to get insight into the underlying molecular phenomena, we decided to use peptides adopting well-defined beta-sheet structures on hydrophobic surfaces. Such peptides were described by De Grado and Lear¹⁶ and consist of alternating leucine (L) and lysine (K) residues to create a hydrophobic repeat, matching a beta-sheet. Using circular dichroism (CD) and Fourier-transform infrared spectroscopy (FTIR), they demonstrated that the synthetic peptide LKLKLKL adopted beta-sheet-stabilized monolayers at apolar interfaces, the hydrophobic leucine residues segregating toward the hydrophobic surface, and the lysine residues toward the water. On the basis of these findings, we reasoned that LK-repeat peptides of variable length were ideal tools to investigate the formation of nuclei, presenting an aggregation-triggering, surface-adsorbed beta-sheet structure to incoming HI molecules.

Here we present a thorough study of the mechanism of formation of HI aggregation nuclei on hydrophobic surfaces, using a series of (LK)nL peptides with varying length and a fluorescent tag. We investigate the dual concentration-dependent effect that these peptides have on HI aggregation kinetics. We demonstrate that their accelerating effect is based on their cooperative and stable adsorption properties, increasing the efficiency of formation of surface-bound prenuclei and that their inhibitory effect is due to an excess of peptide in solution.

## EXPERIMENTAL AND THEORETICAL METHODS

**Human Insulin.** Recombinant human insulin (HI) produced in *Saccharomyces cerevisiae* was purchased from Sigma-Aldrich (ref: I2643). HI solutions TMN buffer (NaCl 125 mM, MgCl₂ 2 mM, Tris-HCl 25 mM, pH 7.4). In order to solubilize insulin, the pH of the solution was lowered to 3.2 using 1 M HCl then adjusted to pH 7.4 with 1 M NaOH using a pH electrode. The HI concentration was then precisely adjusted to 86 µM, measuring the 280 nm absorbance [$ε_{280}$(HI) = 5.53 $mM^{-1} cm^{-1}$)]. The resistivity of the pure water used was 18 MOhm cm at 25 °C. Once adjusted at the proper concentration, the solution was filtered through a 0.22 µm Millex-GV filter unit (ref: SLGV033SS). The solutions were stored at T = 4 °C for less than 2 weeks and refiltered (if not fresh) before use.

**Thioflavin T.** When linked to amyloid fibers, Thioflavin T (ThT) undergoes a fluorescence shift from ($λ_{ex}$ = 342 nm; $λ_{em}$ = 430 nm) to ($λ_{ex}$ = 450 nm; $λ_{em}$ = 482 nm). This property allows the direct quantification of amyloid fibers by fluorescence since the fluorescence of linked ThT is proportional to the amount of amyloid fibers. Thioflavin T (from Sigma-Aldrich, ref: T3516) was solubilized in TMN buffer.

Then the ThT concentration was adjusted to 1 mM by absorbance measurement (in ethanol $ε_{416}$(ThT) = 26.6 $mM^{-1} cm^{-1}$). Finally, solutions were filtered through a 0.22 µm Millex-GV filter (ref: SLGV033SS) and conserved protected from light at T = 4 °C for several months.

**Peptides.** Chemically synthesized (LK)nL peptides as well as the tetramethyl rhodamine-labeled (LK)SL peptide (TAMRA-LK11) (Table 1) were ordered from Genecust

<table><caption>**Table 1. (LK)nL Peptides**</caption><thead><tr><td>peptide name</td><td>peptide sequence</td></tr></thead><tbody><tr><td>LK5</td><td>LKLKL</td></tr><tr><td>LK7</td><td>LKLKLKL</td></tr><tr><td>LK9</td><td>LKLKLKLKL</td></tr><tr><td>LK11</td><td>LKLKLKLKLKL</td></tr><tr><td>TAMRA-LK11</td><td>TAMRA<sub>L</sub>LKLKLKLKLKL</td></tr></tbody></table>

(Luxembourg), solubilized in 10 mM NaOH to a concentration between 75 and 115 µM then filtered through a 0.22 µm filter (Millex-GV ref: SLGV033SS) and stored at T = -20 °C. The concentration of unlabeled peptides was determined using the Quanti Pro Bicinchoninic acid assay (Sigma-Aldrich ref QPBCA). The concentration of TAMRA-LK11 was determined using the molar extinction coefficient of tetramethyl rhodamine, $ε$ = 65 200 $mM^{-1} cm^{-1}$.¹⁷ A linear correlation curve linking fluorescence at 585 nm to the amount of TAMRA-LK11 peptide was established (data not shown).

**Quantification of TAMRA-LK11 Adsorption on Hydrophobic Surfaces.** TAMRA-LK11 was incubated at different concentrations (up to 2.8 µM) in hydrophobic 96-well plates (in triplicate) and shaken at 1200 rpm for 10 min. The solution was then discarded and the well washed with 200 µL of filtered TMN buffer. After discarding the buffer, the adsorbed peptide was desorbed using 0.5% SDS in TMN buffer (1 min at 1200 rpm and 37 °C). The peptide TAMRA-LK11 was then quantified using a calibration curve (see above).

**Measurement of Insulin Aggregation Kinetics.** Aggregation kinetics were done in a 200 µL reaction volume in hydrophobic black 96-well plates (Nunc TM ref: 237105). In each well, aggregation assays were performed in the following conditions: [HI] = 86 µM, [ThT] = 20 µM, [LK peptide] = 0-5 µM, at T = 37 °C and agitation of 1200 rpm on a plate agitator (Heidolph instrument, Titramax 100). The amount of amyloid fibers has been monitored by fluorescence measurement of ThT bound to amyloid fibers ($λ_{ex}$ = 450 nm, $λ_{em}$ = 482 nm). The plates were removed from the incubator/agitator in order to read their fluorescence in a Tecan infinite M1000 plate reader, every 15 min. The bandwidth of the monochromators at the excitation and emission wavelengths was 5 nm. Data were obtained with three independent experiments using five replicates for each experimental condition. Lag times and growth rates were obtained by fitting data points of individual replicates.¹⁴ These were then used to calculate mean values and standard deviations.

**Cooperativity Calculations.** To calculate the Hill coefficient of the cooperative effect of the peptides on the lag time, the experimental data points of the lag times ($τ$) as a function of peptide concentration (see also Figure 1) were fitted to the following equation:

$$τ = τ_{max} - (τ_{max} - τ_{min}) * \frac{[LK]^{n}}{[LK]^{n} + [LK]_{c}^{n}} \quad (1)$$
<figure><img src="image_83.png" /><figcaption>Figure 1. Different effects of LK peptides on the lag time of fibrillation. HI and ThT have been mixed with LK5 (A, ○), LK7 (A, ●), LK9 (B, □) and LK11 (B, ■) peptides at different concentrations and HI fibrillation has been monitored by ThT fluorescence. The solid line represents the fit of the experimental data with the Hill equation (see Experimental and Theoretical Methods). (C) The Hill coefficient (circles) and the critical concentration (squares), corresponding to the one at which 50% of the reduction of the maximal lag time was obtained, are plotted versus the number of LK amino acids. For easier reading, symbols are joined by a hand-drawn dashed line.</figcaption></figure>

where $τ_{max}$ represents the lag time (min) without peptide, $τ_{min}$ the minimal lag time (min) obtained for a given peptide, [LK] the peptide concentration (µM), [LK]c the critical peptide concentration, corresponding to the concentration at which the lag time is half maximal and n the Hill coefficient. The fit was obtained by minimizing the quadratic error between experimental and calculated $τ$, weighted by the standard deviation of the lag time.

For the cooperative adsorption of TAMRA-LK11 on the hydrophobic surface of 96-well plates, the following equation was used to fit the experimental data:

$$[LK11]_{ad} = [LK11]_{max} * \frac{[LK11]_{sol}^{n}}{[LK11]_{sol}^{n} + [LK11]_{c}^{n}} \quad (2)$$

where [LK11]ad represents the concentration (in µM) of the TAMRA-LK11 peptide adsorbed on the well walls, [LK11]sol represents the concentration (in µM) of the TAMRA-LK11 peptide remaining in solution (i.e., [LK11]sol = [LK11]initial - [LK11]ad), and [LK11]max represents the equivalent maximum concentration (in µM) of TAMRA-LK11 peptide adsorbed on the well surface for a given initial TAMRA-LK11 concentration, [LK11]initial, [LK11]c represents the critical [LK11] concentration (in µM), corresponding to the half-maximal concentration of adsorbed TAMRA-LK11, and n represents the Hill coefficient of cooperative peptide surface adsorption. The fit was obtained by minimizing the quadratic error between experimental and calculated peptide concentration, weighted by the standard deviation of the measured concentrations.

**Atomic Force Microscopy.** Glass Functionalization. Borosilicate coverslip glasses ($φ$ = 32 mm, from VWR international ref: 631-0162) were washed successively in SDS, ethanol, water, dried, and then activated in oxygen plasma for 4 min, and treated ($t$ = 2 h) with dimethyldichlorosilane 5% (v/v) in toluene. This process covalently couples the silane to the glass by reacting with exposed Si-OH groups on the glass. The coverslips were then washed in toluene and absolute ethanol, and incubated at $T$ = 100 °C (2 h) (curing) in order to cross-link silane oxides. Functionalization of the glass was controlled by water contact angle analysis (20 µL droplets of pure H₂O). The resulting glass presented a contact angle of 100° ± 5°, indicating a significant hydrophobicity.

**Sample Preparation for the Observation of Insulin Aggregation Nuclei by AFM.** Functionalized borosilicate coverslips were incubated in 6-well NucIon Delta surface plates (Thermo scientific, ref: 140675) under the following conditions: $T$ = 37 °C, 1200 rpm, [HI] = 43 µM, [LK9] = 0 or 0.2 µM, [ThT] = 20 µM in TN (NaCl 125 mM, Tris-HCl 25 mM, pH 7.4). The aggregation state was monitored using fluorescence measurements with a Tecan infinite M1000 apparatus ($λ_{ex}$ = 450 nm; $λ_{em}$ = 482 nm for bound ThT). Before AFM imaging, the samples were washed briefly by dipping in 0.1 µm-filtered TN buffer and dried. Mg was omitted from the buffer to avoid crystal formation during drying.

**Visualization of Insulin Amyloid Aggregation Nuclei by Atomic Force Microscopy.** Samples were imaged in TN buffer using a Dimension Icon (Bruker SAS, Palaiseau, France), equipped with a 150 µm scanner in the intermittent-contact mode. We used a Scanasyst-air Si₃N₄ cantilever (Bruker) with a resonance frequency of 70 kHz and a spring constant of 0.4 N m⁻¹. Images were obtained in the center of the coverslips, where nuclei density was highest. We show representative 2 × 2 µm² topology images. The analysis of the topologies was performed using the NanoScope Analysis software (Bruker).

**Fluorescence Microscopy.** Sample Preparation for the Observation of Insulin Amyloid Aggregates by Fluorescence Microscopy. Fluorescence microscopy observations of HI aggregates were conducted in transparent polystyrene flat-bottom 96 well plates (Greiner bio-one, ref: 655101). The aggregation procedure was the same as for the kinetic measurements. At selected time points, the plate was removed