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Clémence Gruget
To cite this version:
Clémence Gruget. Energy and conformational landscape of Synaptotagmin-1 interacting with mem- branes. Human health and pathology. Université Paris sciences et lettres, 2018. English. �NNT : 2018PSLEE031�. �tel-02128333�
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!
COMPOSITION DU JURY!:
M. RIEU Jean-Paul
Université Lyon-1, Rapporteur!
M. SOELLNER Thomas
Heidelberg University , Rapporteur!
Mme. CRIBIER Sophie UPMC, Présidente du jury Mme. DU ROURE Olivia ESPCI, Examinatrice Mme. PAQUET Claire
Université Paris Diderot, Examinatrice M. ROTHMAN James
Yale University, Examinateur M. PINCET Frédéric
ENS, Directeur de thèse
M. DONALDSON Stephen H Jr ENS, Invité
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Clémence GRUGET soutenue le 11 juin 2018
h
THÈSE DE DOCTORAT
de l’Université de recherche Paris Sciences et Lettres!
PSL Research University
Préparée à l’Ecole Normale Supérieure
Dirigée par Frédéric PINCET
h
Ecole doctorale n°
564 Physique en Ile de FranceSpécialité
BiophysiqueEnergy and conformational landscape of Synaptotagmin-1 interacting with membranes
Paysages energétique et conformationnel d’interaction de la Synaptotagmin-1 avec des membranes
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Remerciements
Je tiens tout d’abord à remercier l’ensemble des membres de mon jury qui ont généreusement accepté de consacrer une partie de leur temps si précieux à l’évaluation de cette thèse. Merci à mes deux rapporteurs, Thomas Söellner et Jean-Paul Rieu d’avoir rédigé leurs rapports en un temps record, ainsi qu’à mes examinateurs, Sophie Cribier, Olivia Du Roure, Claire Paquet et James Rothman pour leurs questions témoignant de l’intérêt porté à mon travail.
Je ne pourrai jamais assez remercier mon directeur de thèse Frédéric Pincet de m’avoir prise sous son aile avec un grand sourire il y a quatre ans lorsque j’ai frappé à la porte de son laboratoire à la recherche d’une thèse atypique. La confiance totale qu’il m’a accordée, déconcertante par moment, a été un formidable outil d’apprentissage de l’autonomie, de la patience et de la persévérance. En plus de l’inépuisable source de connaissances qu’il a partagée avec moi autant que possible, ce sont aussi ses qualités humaines qui m’auront fait grandir bien plus que j’aurais pu le soupçonner.
J’ai eu la chance inouïe de toujours avoir pu compter sur un autre superviseur non moins talentueux, Steve Donaldson. Ce n’est pas exagéré de ma part que de dire que sans sa présence constante à mes côtés, dans les bons moments et surtout dans les instants de déconvenues et de doutes, je n’aurais pas pu relever ce défi, le poids de l’échec ayant été trop lourd à porter sur mes seules épaules. Son expertise mais surtout sa bienveillance, son calme olympien, sa patience presque surnaturelle face aux épreuves qui se sont accumulées, son optimisme, sa profonde foi en la science et en nos travaux ont été la lumière avec laquelle bien souvent je me suis éclairée.
J’ai également pu trouver un ange gardien en la personne de Jim Wang. Nos chemins se sont croisés dès le premier jour de ma thèse, lors du passage de relais autour du SFA. Je garderai toujours une profonde admiration pour ce modèle d’abnégation au service de la recherche qu’il représente à mes yeux. En plus de son talent, sa bonté et sa gentillesse m’ont apporté bien plus que ce dont il a conscience. Même si nos parcours respectifs nous ont séparé assez rapidement, je suis certaine que notre amitié perdurera.
Je serai éternellement reconnaissante au professeur James Rothman de m’avoir intégrée avec enthousiasme et bienveillance aux travaux de sa fabuleuse équipe, m’accueillant dans ses laboratoires, et m’offrant ainsi un cadre d’apprentissage exceptionnel. Pouvoir partager avec lui mes avancées souvent laborieuses, auxquelles il a toujours répondu avec patience et encouragements, fut un privilège.
A toute son équipe, thésards, post-docs, techniciens, assistants, à Yale et à UCL, eux qui ont été ma deuxième famille pendant mes séjours d’expatriation, à la fois humaine et scientifique, je dois également beaucoup. Même lorsqu’eux même traversaient des moments difficiles, leur gentillesse et leur soutien ont été sans limite. Jamais je ne me suis vue reprocher le temps pourtant considérable que j’ai pu leur faire perdre en questions et demande d’aide en tout genre, sollicitations auxquels ils répondaient toujours avec un sourire. J’espère pouvoir un jour leur rendre autant qu’ils m’ont donné.
Merci à mes mentors des premières heures, Isabelle, Claire et Benjamin, d’avoir partagé avec moi leurs techniques de survie. Sans eux ma thèse n’aurait certainement pas été la même.
Merci d’avoir été là, dans les bons comme dans les mauvais moments. De ces longues heures passées à vos côtés au sous-sol du département sera née une amitié toute particulière pour moi.
Ce fut un plaisir de partager ces années avec la nouvelle et talentueuse génération du LPS.
Jean Noël Dominique va poursuivre les péripéties du SFA et finira j’en suis sûre par le dompter mieux que personne. Sophie fera naître des entrailles de ses souris de fécondes découvertes. La Rachid’s team, Lucie, Kalthoum, Aymeric, Vincent, et tous les autres, feront de leurs micro manips des macro résultats. Merci à eux, ainsi qu’à Loïc qui m’a régulièrement arrachée à mes machines obscures pour m’aérer l’esprit. Je leur souhaite à tous l’épanouissement et la réussite qu’ils méritent.
Merci également à tous les chercheurs permanents, Christine Gourier, Eric Perez, Lionel Foret, Nicolas Rodriguez, Rachid Thiam, Wladimir Urbach, qui m’ont entourée de près ou de loin toutes ces années, pour répondre à mes questions, partager leur passion et leur savoir, me réconforter ou simplement discuter autour d’un café.
Merci au personnel administratif du LPS, en particulier à mes fidèles alliés, Annie Ribaudeau, Benoît Paulet, et Nora Sadaoui. Merci de n’avoir jamais été à court d’idées pour rattraper mes bêtises, de m’avoir aidée dans mes commandes farfelues et de m’avoir permise de partir si souvent en mission à la dernière minute. Merci pour leur professionnalisme, leur gentillesse et leur patience à toute épreuve. Merci à Zaire Dissi, le roi des ordis, et à tous les autres MacGyver de l’ombre. Merci également à Laura Baron-Ledez et Jean François Allemand de l’ED PIF, ainsi qu’à Stéphane Emery et Krisztina Hever-Joly du bureau des doctorants de l’ENS, qui auront été plus que compréhensifs avec moi… Qu’il est bon de se sentir bien entourée !
Merci bien sûr à tous mes amis m’ayant supportée pendant ces années, mon crew, qui ont su écouter mes angoisses et mes doutes, ont pu me réconforter et me redonner le sourire quand j’en avais le plus besoin. Auprès d’eux je peux difficilement mentir et je n’ai pas toujours montré le meilleur de moi-même. Merci de m’avoir comprise et aimée malgré tout. Un merci tout particulier à mon Manu pour m’avoir remise dans le sens de la marche lorsque j’étais prête à dérailler. Je ne serai pas allée au bout sans eux. Merci à mon cousin David pour son aide matérielle, humaine et intellectuelle miraculeuse. Merci aussi à mes amis du COGE venus fêter ma délivrance avec une belle surprise musicale !
Merci enfin à ma famille de m’avoir suivie dans cette folie les yeux fermés même lorsque je n’étais pas certaine de mon chemin, et de m’avoir entourée de sa confiance, même lorsque je commençais à en manquer. Papa, maman, mes sœurs adorées, je vous aime.
“Anything that can go wrong will go wrong.”
Murphy’s Law
TABLE OF CONTENTS
CHAPTER I: INTRODUCTION AND BACKGROUND 1
I. MOLECULAR ACTORS OF NEUROTRANSMISSION 2
I.1 SNARE/SM PROTEINS: THE DRIVING FORCE OF THE FUSION MACHINERY 3
I.2 COMPLEXIN 5
I.3 SYNAPTOTAGMIN: THE CALCIUM SENSOR OF NEUROTRANSMISSION 5
II. MOLECULAR ANATOMY OF SYNAPTOTAGMIN-‐1 7
III. EXISTING HYPOTHESIS ON THE ACTION OF SYNAPTOTAGMIN-‐1 10 III.1 SYNAPTOTAGMIN-‐1 ACTS AS A CLAMP THAT IS RELEASED WITH CALCIUM 11 III.2 SYNAPTOTAGMIN-‐1 REMOVES THE COMPLEXIN CLAMP UPON CALCIUM TRIGGER 12 III.3 SYNAPTOTAGMIN-‐1 REGULATES THE GAP BETWEEN THE PLASMA MEMBRANE AND THE SYNAPTIC VESICLE
IN A CALCIUM DEPENDENT MANNER 14
III.4 SYNAPTOTAGMIN-‐1 PROMOTES MEMBRANE FUSION BY INDUCING CURVATURE 14 IV. PREVIOUS MEASUREMENTS ON SYNAPTOTAGMIN-‐1 MEMBRANE INTERACTIONS 15
V. AIMS OF THIS PHD 16
CHAPTER II: THE SURFACE FORCE APPARATUS: GENERAL PRINCIPLES AND APPLICATION TO THE MEASUREMENT OF SYNAPTOTAGMIN-‐1 INTERACTIONS WITH MEMBRANES 19
I. THE SURFACE FORCE APPARATUS SET UP 20
I.1 GENERAL PRESENTATION OF THE SFA 20
I.2 MICA SURFACES PREPARATION 21
I.3 SURFACES DISPLACEMENT SYSTEM 21
I.4 DISTANCE MEASUREMENT PRINCIPLE 22
I.5 FORCE MEASUREMENT PRINCIPLE 24
I.6 INTERACTION ENERGY BETWEEN THE SURFACES 25
I.6.1 Relation between the force, the distance and the interaction energy 25
I.6.2 Description of a typical SFA force run 25
II. FUNCTIONALIZATION OF THE MICA SURFACES WITH A LIPID BILAYER 26 II.1 THE LANGMUIR BALANCE AND LANGMUIR BLODGETT DEPOSITION PRINCIPLE 27
II.2 ISOTHERMAL COMPRESSIONS 29
III. PRELIMINARY TESTS OF THE SFA ON MICA-‐MICA AND DMPE-‐DMPE FORCE RUNS 34
III.1 MICA-‐MICA FORCE PROFILE 34
III.2 DMPE-‐DMPE FORCE PROFILE 37 IV. DESIGN, PURIFICATION AND MEMBRANE BINDING TESTS OF SYNAPTOTAGMIN-‐1 39 IV.1 EXPRESSION AND PURIFICATION OF SYNAPTOTAGMIN-‐1 39
IV.1.1 DNA design 39
IV.1.2 Purification of Synaptotagmin-‐1 41
IV.2 MEMBRANE BINDING ASSAYS 43
IV.2.1 Liposome binding assays 43
IV.2.2 Syt1 binding to PS/PIP2/PC liposomes assay 44
IV.2.3 Syt1 binding to DGS-‐NTA(Ni)/PC liposomes assay 46
IV.2.4 Syt1 binding to GUVs 48
IV.2.5 Binding of Syt1 to supported bilayers 49
V. CONCLUSION 53
CHAPTER III: DIRECT MEASUREMENT OF THE BINDING ENERGY OF SYNAPTOTAGMIN-‐1
TO AN ANIONIC MEMBRANE 55
I. EXPERIMENTAL SET UP FOR THE SFA: COMPARISON WITH A PHYSIOLOGICAL SYNAPTIC VESICLE -‐ PLASMA
MEMBRANE INTERFACE 55
I.1 MEMBRANE COMPOSITION: REDUCED COMPLEXITY FOR MORE SPECIFIC INTERACTIONS 56 I.2 SYNAPTOTAGMIN-‐1 INCORPORATION TO THE CIS-‐MEMBRANE: HIS-‐TAG VERSUS MEMBRANE INSERTION 59
I.3 MOLECULAR CROWDING 59
I.4 CONFINEMENT DIMENSIONS 61
I.5 MEMBRANE DEFORMABILITY 62
II. RESULTS 62
II.1 EFFECT OF ANIONIC LIPIDS ON SYT1 INTERACTIONS WITH LIPID MEMBRANES 63 II.1.1 Approach: extraction of the protein surface density Γ and radius of gyration Rg from a
polymer mushroom model 64
II.1.2 Separation: measurement of Syt1 binding energy to an anionic membrane in EGTA 65 II.2 EFFECT OF DIVALENT CATIONS (CA2+ AND MG2+) ON SYT1 ADHESION TO MEMBRANES 67 II.2.1 Approach: the Syt1-‐membrane repulsive force profile is not affected by the presence of
divalent cations 68
II.2.2 Separation: measurement of Syt1 binding energy to an anionic membrane in Mg2+ and Ca2+
68
II.2.2.1 The binding energy increases with Mg2+ and Ca2+ 68 II.2.2.2 A molecular rearrangement occurs during contact time and is correlated with the
increase in binding energy 69
II.2.2.3 Bond lifetime estimation 72
III. DISCUSSION 74 III.1 CALCIUM INDEPENDENT BINDING PROPERTIES OF SYT1 74 III.2 POTENTIAL MECHANISM FOR THE SYT1-‐MEMBRANE REARRANGEMENT LEADING TO THE INCREASED BINDING ENERGY MEASURED IN THE PRESENCE OF MG2+ AND CA2+ 76
IV. CONCLUSION 79
CHAPTER IV: MUTAGENESIS STUDY OF SYNAPTOTAGMIN-‐1–MEMBRANE INTERACTIONS
81
I. MUTAGENESIS DESIGN 81
II. RESULTS: SFA STUDY ON THE FOUR SYT1 MUTANTS 86
II.1 STUDY OF THE KAKA MUTANT 87
II.2 STUDY OF THE KAKAC2AB MUTANT 91
II.3 STUDY OF THE C2AB MUTANT 93
II.4 STUDY OF THE KAKAC2AB MUTANT 95
III. DISCUSSION 98
III.1 CALCIUM INDEPENDENT MEMBRANE BINDING ENERGY OF SYT1 100 III.2 CALCIUM DEPENDENT MEMBRANE BINDING ENERGIES OF SYT1 100 III.2.1 Estimation of the global calcium dependent membrane binding energy 100
III.2.2 Energies of individual C2 domains 101
III.2.2.1 Calcium dependent membrane binding energy estimation of the C2A domain 101 III.2.2.2 Calcium dependent membrane binding energy estimation of the C2B domain 102 III.2.3 Models of the energetic landscape of Syt1-‐membrane interaction in calcium 103
III.2.3.1 Preliminary remarks 103
III.2.3.2 Additive models for the membrane binding energies of C2A and C2B domains 105 III.2.3.3 Cooperative model for the membrane binding energies of C2A and C2B domains 106
IV. CONCLUSIONS AND PERSPECTIVES 112
CONCLUSION AND OUTLOOKS 115
APPENDICES 119
ANNEX A: LIPOSOME PREPARATION 119
ANNEX B: GIANT UNILAMELLAR VESICLES PREPARATION BY ELECTROFORMATION 121
ANNEX C: PUBLICATION I 123
ANNEX D: PUBLICATION II 135
BIBLIOGRAPHY 145
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Chapter I: Introduction and background
Neural communication underlies all brain activity. It governs our thoughts, feelings, sensations, and actions. Communication between two neurons occurs at specialized cell junctions called synapses, where two communicating neurons are separated by a synaptic cleft. The presynaptic neuron releases chemical, known as neurotransmitters, into the synaptic cleft where neurotransmitters bind to receptors on the surface of the postsynaptic neuron, hence propagating the signal. This release of neurotransmitters requires the fusion of the synaptic vesicles encapsulating neurotransmitters with the plasma membrane of the neuron in response to an action potential, also known as nerve impulse, within the sending neuron.
A hallmark of synaptic transmission is its speed, and synchronicity, without which a coherent cognitive activity would be impossible. In contrast to most intracellular membrane fusion processes, synaptic vesicle fusion occurs in less than one millisecond timescale, making it the fastest membrane fusion event in the cell (Thomas C. Südhof 2013). This requires the synaptic vesicles to be positioned at the right location and ready for fusion upon calcium entry in the neuron. To reach this “primed” state most of the synaptic vesicles undergo a series of maturation steps, illustrated in figure I.1. Once filled with neurotransmitters, synaptic vesicles are recruited or “tethered” to specialized sites at the presynaptic plasma membrane called active zones. There, the vesicles attach closely or “dock” to the fusion sites. Docked vesicles finally undergo a “priming” step, that renders them ready for fast membrane fusion (Jahn &
Fasshauer 2012). We will describe below the current view of the molecular arrangements in the primed state. When an action potential invades a presynaptic nerve cell, voltage-gated calcium channels localized in the active zone open briefly, resulting in a sharp rise of the calcium concentration in the vicinity of the docked vesicles. This calcium influx is the chemical trigger for the primed vesicles to fuse with the membrane of the neuron, leading to the fast and synchronous release (also called “evoked” release) of their content in the synaptic cleft, and to the propagation of the signal to the postsynaptic neuron.
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Figure I.1: Neuronal trafficking overview. Synaptic vesicles are filled with neurotransmitters and stored in the cytoplasm of the nerve terminal. The loaded synaptic vesicles are then targeted to specialized sites of release called the active zone where the vesicles dock. They then undergo priming steps that make them competent to fuse. Upon the arrival of an action potential, the resulting influx of calcium ions through voltage-activated Ca2+
channels trigger exocytosis. After fusion, the vesicle is retrieved by endocytosis, mediated by the formation of a clathrin-coated vesicles. After clathrin uncoating, synaptic vesicles are regenerated within the nerve terminal and reloaded for another round of exocytosis. Extracted from (Jahn & Fasshauer 2012)
The release of the synaptic vesicles from the primed state therefore requires a calcium sensor.
This calcium sensor has already been identified as a protein called Synaptotagmin. The purpose of this PhD is to elucidate the interplay between Synaptotagmin, calcium and membranes. In this introduction we will present the various molecular players and provide a comprehensive description of Synaptotagmin and its putative mode of action.
I. Molecular actors of neurotransmission
How do the synaptic vesicle and the plasma membrane fuse during transmitter release? How does calcium trigger synaptic vesicle fusion? The molecular mechanism that permits such controlled release has been the subject of intense research (reviewed in (Jahn & Fasshauer 2012; Rizo & Xu 2015; Chapman 2008). Several key players are proteins located either in the membrane of the synaptic vesicle, the plasma membrane of the neuron or in the cytoplasm (figure I.2).
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SNARE (Soluble NSF(N-ethylmaleimide-sensitive factor) Attachment Protein Receptor) proteins along with SM (Sec1/Munc18-like) proteins (figure I.2 A and B) are the universal fusion machinery for different intracellular fusion process. In the case of neurotransmission, the need for a fast and calcium triggered fusion is governed by additional proteins that have emerged as key regulators: Synaptotagmin and Complexin (figure I.2 C and D). We will now give a brief description of these proteins.
Figure I.2: Structure of the main proteins involved in the fusion process. A: SNARE complex of VAMP/Synaptobrevin (blue helix), SNAP25 (green and yellow helices). B: Munc18. C: Complexin, bound to the SNARE complex (purple). D: Synaptotagmin. Extracted from (Südhof & Rothman 2009)
I.1 SNARE/SM proteins: the driving force of the fusion machinery
The core of the membrane fusion machinery is formed by a set of three membrane proteins called the SNARE proteins, for Soluble NSF (N-ethylmaleimide-sensitive factor) Attachment Protein Receptor (Südhof & Rothman 2009). They fall into two categories: the v-SNAREs in transport vesicles and t-SNAREs in target membranes. The t-SNAREs is an heterodimer made oftwo proteins called SNAP-25 and syntaxin-1 emanating from the presynaptic plasma membrane, and the v-SNARE consists of the vesicle associated membrane protein VAMP2, also called synaptobrevin-2. Individual SNARE proteins are unfolded, but v- and t-SNAREs engage spontaneously to assemble into a very stable four-helix bundle (formed by the conserved SNARE motif) three α-helices emanate from the t-SNAREs (two α-helices are brought by SNAP-25, one by syntaxin-1), and one α-helix emanates from the v-SNARE synaprobrevin-2 (Söllner et al. 1993). This assembly that forms between membranes is called a “trans-SNARE complex”, also known as a “SNAREpin”. This complex is the force
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generator of membrane fusion: the zippering of the SNAREs from their membrane-distal N- terminal domain to their membrane proximal C-terminal domain, releases about 35 kBT of energy per SNAREpin (Li, Pincet, Perez, William S Eng, et al. 2007) which is translated into a force that pulls the apposing bilayers close together and fuses them (figure I.3). The activation energy required for the fusion of lipid bilayers is estimated to be between 20 to 150 kBT, depending on the model, and a recent bulk lipid-mixing assay measured an activation energy of about 30 kBT (François-Martin et al. 2017), a barrier that could be quickly overcome by the SNARE zippering. In fact, one SNARE pinmay be sufficient for membrane fusion in vitro (Van Den Bogaart et al. 2010), although probably not at the physiological rate (Xu et al. 2016) for which several SNAREs are likely to be needed to act in concert.
Figure I.3: Illustration of the SNARE-mediated fusion. Three helices anchored in one membrane (the t-SNARE) assemble with the fourth helix anchored in the other membrane (v-SNARE) to form trans-SNARE complexes, or SNAREpins. Assembly proceeds progressively from the membrane-distal N termini toward the membrane- proximal C termini of the SNAREs. This generates a force that pulls the bilayers together, forcing them to fuse.
In a physiological context, the SNAREs need to be made competent for fusion. This is achieved by additional proteins such as the SM protein Munc18 and Munc13. They are involved in setting up the SNAREs for assembly and perhaps in guiding them through the initial part of the assembly pathway, but it is still not understood how exactly they operate (Jahn & Fasshauer 2012).
SNARE proteins generally take 0.1 to 1 second to spontaneously complete fusion (Weber et al. 1998). How can neurotransmitters be released 1000 to 10 000 times faster by the same SNAREs? Two proteins have been shown to be necessary and sufficient, in addition with the SNARE proteins, for synaptic transmission: Complexin and Synaptotagmin-1.
Membrane(fusion(
t/SNARE(
v/SNARE(
Vesicule(
Target(membrane(
5
I.2 Complexin
Complexin is a small, soluble protein that binds tightly to the SNARE four helices bundle when it is in a partially zippered state via a central helix (figure I.2C). Removal of Complexin results in an increase in the so-called spontaneous release, which is a type of release that is not triggered by an action potential. This suggests that Complexin has an important role in clamping synaptic transmission and preventing the spontaneous release of neurotransmitters.
Deletion of Complexin also reduces the amount of evoked release, suggesting that Complexin also facilitates fusion. The release occurs with longer and variable delay after the action potential (the release becomes asynchronous). Hence Complexin is able to both activates and clamps neurotransmitter release, presumably by independent different mechanisms (Martin et al. 2011). Such dual inhibitory and stimulatory role for fusion is not completely understood (reviewed in (Mohrmann et al. 2015).
If Complexin is essential for synchronous release, it cannot alone account for the full regulation of the calcium-triggered neurotransmission process, as Complexin does not bind calcium. Another protein plays the role of the calcium sensor: Synaptotagmin.
I.3 Synaptotagmin: the calcium sensor of neurotransmission
Calcium-triggered ultra fast synaptic vesicle fusion allowing the release of neurotransmitters was first revealed by Bernard Katz in the 1960’s (Katz & Miledi 1969). Since then, scientists have been looking for the calcium sensor of neurotransmission. This quest was largely completed by the research of Südhof and colleagues that worked over two decades to show that calcium dependent exocytosis is mediated by proteins called Synaptotagmins.
Synaptotagmins are evolutionarily conserved transmembrane proteins. There are 15 isoforms of mammalian Synaptotagmins, localized to synaptic and secretory vesicles or the plasma membrane. All Synaptotagmins are composed of an N-terminal transmembrane region, a linker of variable length and two calcium-binding modules, the C2 domains, termed C2A and C2B (figure I.4). The C2 domain is a conserved sequence motif of 130–140 amino acid residues that share a common overall fold: an eight-stranded anti-parallel β-sandwich
6
consisting of a pair of four-stranded β-sheets, involved in calcium and phospholipid binding(Corbalan-Garcia & Gómez-Fernández 2014).
Figure I.4: Domain structures of mouse Synaptotagmins. All Synaptotagmins are composed of an N-terminal transmembrane region, a linker of variable length and two functional C2 domains. Extracted from (Gustavsson
& Han 2009)
The first isoform of Synaptotagmin that was identified as a calcium sensor for neurotransmission was Synaptotagmin-1. Südhof and colleagues performed electrophysiological analyses on Synaptotagmin-1 knockout mice, and discovered that the fast synchronous transmitter release was ablated, establishing that Synaptotagmin-1 is essential for fast calcium-triggered release (figure I.5) (Geppert et al. 1994). Interestingly, the slow asynchronous release was not affected by the Synaptotamgin-1 knockout, indicating that the role of Synaptotagmin-1 is specific to the synchronous calcium-triggered release. They further showed that this essential role of Synaptotamin-1 was directly related to its calcium binding properties. Introducing a point mutation in the endogenous Synaptotagmin-1 mouse gene that decreased the Synaptotagmin-1 calcium binding affinity by about two fold, also causes a 2-3 fold decrease in the calcium sensitivity of release measured by electrophysiology (Fernández- Chacón et al. 2002). Conversely, a gain of function study showed that mutations in the C2 domains of Synaptotagmin-1 that increased its apparent calcium affinity was correlated to an increase in the calcium sensitivity of release (Rhee et al. 2005). Hence, the calcium affinity of Synaptotagmin-1 determines the calcium affinity of synaptic exocytosis. This formally established Synaptotagmin-1 as the calcium dependent switch to activate fusion.
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Figure I.5: Physiological evidence establishing Synaptotagmin-1 as the calcium sensor of neuronal exocytosis:
Synaptotagmin-1 knockout ablates fast synchronous neurotransmitter release. Traces showing recordings from neurons cultured from wild-type (a) and Synaptotagmin-1 knockout (b) mice. Traces depict synaptic responses to isolated action potentials; standard (top) and expanded views (bottom) illustrate that the Synaptotagmin-1 knockout completely ablates fast synchronous release and increases slow asynchronous responses. Extracted from (Thomas C Südhof 2013).
Not all Synaptotagmin C2 domains bind to calcium. In fact, based on sequence similarities and subsequent confirmation by biochemical analyses, only eight Synaptotagmins bind to calcium, namely Synaptotagmin-1, -2, -3, -5, -6, -7, -9 and -10 (reviewed in (Gustavsson &
Han 2009). Among these isoforms, in addition to Synaptotagmin-1, Synaptotagmin-2 and Synaptotagmin-9 are also involved in evoked synchronous release, at different types of synapses. Synaptotagmin-7 has been shown to be important for the slower asynchronous release (Bacaj et al. 2013). The inability of other Synaptotagmins to bind calcium is explained for the majority by the lack of critical residues involved in calcium binding.
Synaptotagmin-1 is the most important isoform for evoked released, and will be the focus of our work.
II. Molecular anatomy of Synaptotagmin-‐1
Synaptotamin-1 is a 65 kDa protein associated with the synaptic vesicle (Takamori et al.
2006) via its N-terminal transmembrane helix. A 61-residue linker connects the
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transmembrane domain and the tandem C2 domains, C2A and C2B, which are separated with a 9-residue flexible linker domain. The main properties of Synaptotagmin-1 arise from its C2 domains, which confer the protein distinct calcium independent and calcium dependent functions, as expected from its role as a calcium sensor of evoked release.
Figure I.6: Ribbon diagram of the crystal structure of Synaptotagmin-1 cytoplasmic domain (structure PDB 3R83) where we highlighted the main calcium-independent (in red) and calcium-dependent (calcium residues in yellow and hydrophobic residues in purple). In the absence of calcium, Synaptotagmin-1 binds the negatively charged lipids (PIP2 and PS) localized in the plasma membrane, presumably docking the vesicle in the active zone, through a patch of four consecutive lysine residues (K324/325/326/327).
Synaptotagmin-1 has been shown to bind to the membranes in the absence of calcium. A stretch of four consecutive lysine residues (K324–K327), also referred to as a “polylysine patch”, located at the side of the Synaptotagmin-1 C2B domain, has been proposed to bind to the negatively charged phosphatidylinositol (4,5)-biphosphate (PIP2) lipid clustered on the plasma membrane (Van Den Bogaart et al. 2011; Bai et al. 2004; Lai et al. 2015; Park et al.
2012) (figure I.6). This interaction could be important for the initial stage of docking of the synaptic vesicles to the plasma membrane of the neuron. The polylysine patch/PIP2 interaction will be characterized in this work.
Calcium independent binding site (polybasic patch)
Linker'region' Transmembrane' domain'
421'
C2B' C2A'
C6term' N6term'
143' 82'
Plasma&membrane&
Anionic lipids (PIP2, PS)
+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&+&&&
Hydrophobic loop Calcium binding site Synaptotagmin61'
Synap/c&vesicle&
9
The C2 domains are composed of a stable eigth-stranded β-sandwich with flexible loops emerging from the top and bottom (figure I.7). Two distinct loops per C2 domains, termed loop 1 and loop 3, form a negatively charged pocket that can coordinate calcium ions (Striegel et al. 2012; Herrick et al. 2009). Synaptotagmin-1 C2A domain ligates three calcium ions via five asparate and one serine residues (D172, D178, D230, D232, D238, S235) while the C2B lacks the binding site for calcium 3 and binds only two calcium ions via four asparate residues (D303, D309, D363, D365, D371) (figure I.7). In both C2A and C2B, coordination of calcium ions is incomplete. This allows for a calcium dependent binding of Synaptotagmin-1 to a ternary component that can complete the calcium coordination sphere, such as phospholipids.
The affinity of Synaptotagmin-1 for calcium is indeed strongly enhanced by the presence of the anionic lipids phosphatidylserine (PS) and phosphatidylinosotinol (PIP2) (Van Bogaart et al. 2012).
Non-polar residues (also referred to as hydrophobic loops), are present nearby the calcium binding sites, at the tips of both C2A and C2B calcium binding loops (F231/F234/S235 and V304/Y364/I367 in the C2A and C2B domains respectively). The negatively charged residues and the negative lipids repel each other which forces the hydrophobic loops to be away from the membrane thereby impeding them from inserting in the membrane. Calcium binding to the negatively charged pocket effectively neutralizes electrostatic repulsion between the binding site and the membrane, which allows both hydrophobic loops 1 and 3 to insert into the membrane (Herrick et al. 2006; Chapman & Davis 1998). Electron Paramagnetic Resonance (EPR) studies showed that hydrophobic loop 1 and 3 of both C2A and C2B penetrate an anionic bilayer below the level of the lipid phosphates at a depth of 1 nm (Herrick et al. 2006) (figure I.8).
This “electrostatic switch” between the calcium-unbound and calcium-bound state is at the core of Synaptotagmin-1 function, serving as the power stroke to activate fusion. This switch will also be studied in the course of our measurements.
10
Figure I.7 : Structure of Synaptotamin-1 C2A (orange) and C2B (blue) calcium binding loops, termed loop 1 and loop 3. The C2A domain ligates three calcium ions via five asparate and one serine residues, while the C2B lacks the binding site for calcium 3 and bind only two calcium ions. Extracted from (Rizo et al. 2006).
Figure I.8 : Model for the position and docking of Synaptotagmin C2 domains at an anionic membrane interface obtained by Electron Paramagnetic Resonance (EPR). Both loops 1 and 3 are inserted into the membrane.
Extracted from (Herrick et al. 2006)
III. Existing hypothesis on the action of Synaptotagmin-‐1
Several partially overlapping mechanisms have been proposed for how Synaptotagmin-1 triggers fusion. The first group of models involves that the SNARE complex is arrested at a partially zippered state because of a “clamp” that one way or another inhibits its full assembly. Synaptotamgin-1 may either serve as the clamp, stopping SNAREs assembly until calcium triggering. Alternatively, Complexin is the clamp and Synaptotamgin-1 binding to
11
calcium results in the removal of Complexin from the SNARE complex. Other models center Synaptotagmin-1 mechanism on its membrane binding action, by regulating the distance between the plasma membrane of the neuron and the synaptic vesicle, and promoting bending work on the membranes.
III.1 Synaptotagmin-‐1 acts as a clamp that is released with calcium
Synaptotagmin-1 acceleration of fusion upon calcium trigger has been proposed to be the result of a “clamping” function of Synaptotagmin-1 preventing the full SNARE complex assembly that would be released upon calcium binding (Chicka et al. 2008).
In line with this idea, recent studies electron microscopy studies showed that the cytosolic domain of Synaptotagmin-1 (C2AB) can oligomerise in a circular arrangement (a “ring-like”
structure) on membrane containing anionic lipids (PS and/or PIP2) (Zanetti et al. 2016; Wang et al. 2014; Rothman et al. 2017). This “ring-like” oligomer disappeared upon addition of calcium. The calcium disruption of the rings requires functional calcium binding of the C2B domain of Synaptotagmin-1, as mutations of the calcium binding site of C2B cancelled the calcium-dependent disassembly of the ring. These observations led to a model where in the absence of calcium, Synaptotagmin-1 binds to anionic membrane through the electrostatic interaction between the polylysine patch located on its C2B domain and the negatively charged PIP2 lipids located in the plasma membrane. Synaptotagmin-1 ring-like oligomers form at the interface between synaptic vesicle and the plasma membrane, acting like a washer.
This would sterically hinder fusion by keeping the membrane apart and preventing the SNARE from completing their full assembly. Upon calcium influx, the C2B calcium binding loops of the C2B domain insert into the membrane, causing the ring to disassemble. The removal of this barrier allows for a fast zippering of the SNAREs and fusion, synchronized to the calcium influx (figure I.9) (Zanetti et al. 2016).
12
Figure I.9: ‘Washer’ model for the regulation of neurotransmitter release by Synaptotagmin-1. A: The synaptic vesicle (SV) docking interaction of the Synaptotagmin-1 polylysine motif (blue dots) with the PIP2 (yellow dots) on the plasma membrane positions the Synaptotagmin-1 on the membrane to promote the ring-oligomer formation. The ring assembly might precede the engagement of the SNARE proteins. B: Synaptotagmin-1 ring- oligomers assembled at the synaptic vesicle-plasma membrane interface act as a spacer or ‘washer’ to separate the two membranes. The height of the ring (~4 nm) would allow the partial assembly of the SNARE complex, but prevent complete zippering and thus, block fusion. C: Upon binding calcium ions (red dots), the Ca2+ loops that locates to the oligomeric interface, reorients and inserts into the membrane, thus disrupting the ring oligomer to trigger fusion and release neurotransmitters. Thereby, the Syt1 ring oligomers will synchronize the release neurotransmitters to the influx of calcium ions. D: Negative-stain electron microscopy analysis of ring-like oligomers formed by the cytoplasmic domain of Synaptotagmin-1 (Syt1CD WT) on PS:PC (1:3) lipid monolayers in a physiological buffer. PC: phosphatidylcholine. PS: phosphatidylserin. Adapted from (Zanetti et al. 2016)
III.2 Synaptotagmin-‐1 removes the Complexin clamp upon calcium trigger
Synaptotagmin-1 is able to bind to the t-SNAREs in the plasma membrane when the complex is maintained in its half zippered state, presumably by Complexin, both in the presence and absence of calcium. This suggests that Synaptotagmin-1 binding to calcium would trigger SNARE complex assembly by inserting into the bilayer along with the attached SNAREpin, breaking the clamping interaction of Complexin (Chicka et al. 2008; Krishnakumar et al.
2013).
A recent crystal structure (Zhou et al. 2015) of the Synaptotagmin-1–SNARE complex showed that this binding occurred through polar interactions between multiple charged
D"
13
residues of the C2B domain of Synaptotagmin-1 and charged residues of both t-SNAREs, SNAP25 and Syntaxin-1 (figure I.10, called the “primary interface”).
Another crystal structure of the primed pre-fusion SNARE–Complexin–Synaptotagmin-1 complex revealed a tripartite interface between Synaptotagmin-1 and both the SNARE complex and Complexin (Zhou et al. 2017). Synaptotagmin-1 contacts both Complexin and the t-SNARE syntaxin-1 through helices located on its C2B domain, forming a six helices bundle structure with four helices coming from the SNARE complex and one helix coming from Complexin central helix (figure I.10 A, C2B orange C2B). Simultaneously, a second Synaptotagmin-1 molecule interacts with the SNARE through polar interactions between multiple charged residues of the C2B domain of Synaptotagmin-1 and both t-SNAREs SNAP25 and Syntaxin-1, called the “primary interface” (figure I.10 A, grey C2B). Neither the primary nor the tripartite interface involved the calcium binding loops of Synaptotagmin-1, implying that the binding is independent of calcium. Hence the authors proposed that before calcium influx, the tripartite binding between the SNAREs, Complexin and Synaptotagmin-1 locks the SNAREs in energized a half zippered state, maintaining it ready for fast calcium- triggered fusion (figure I.10 B). The complex is “primed”. Calcium binding to the loops of Synaptotagmin-1 and their subsequent insertion in the membrane unlocks the tripartite interface, allowing the full zippering of the SNAREs, and calcium-triggered fusion to proceed. Calcium binding to the Synaptotagmin-1 molecule involved in the primary interface and resulting membrane loop insertion would also lead to an “en bloc” displacement of the Synaptotagmin-1-SNARE complex, also helping SNARE-mediated fusion possibly by destabilizing the membranes (figure I.10 C). Interestingly, both interfaces are essential for evoked neurotransmitter release, as shown by its strong decrease when mutations of the residues involved in one or the other interface were introduced (Zhou et al. 2017).
14
Figure I.10: Crystal structure of the Synaptotagmin-1-SNARE-Complexin interface, involving two molecules of Synaptotagmin-1. A : One C2B domain (orange) contact both Complexin (Cpx) and the SNARE helice bundle (tripartite interface), while another C2B domain (grey), interacts with the SNAREs (primary interface). B : Model of a primed Syt1–SNARE–Cpx–Syt1 (‘locked’) complex situated between the synaptic vesicle and plasma membranes. C: Upon unlocking and Ca2+ triggering the SNARE complex fully zippers, and fusion occurs. Adapted from (Zhou et al. 2017)
III.3 Synaptotagmin-‐1 regulates the gap between the plasma membrane and the synaptic vesicle in a calcium dependent manner
Without necessarily interacting with the SNARE complex, Synaptotagmin-1 has been proposed to cooperate with them in inducing membrane fusion just by reducing the distance between the synaptic vesicle membrane and the plasma membrane in a calcium dependent manner (Kuo et al. 2011; Herrick et al. 2009). Insertion of the calcium loops would expose the positively charged residues present on the opposite side of the calcium binding site of C2B to the opposite membrane, promoting the simultaneous binding of both membranes by C2B, resulting in their close apposition (Xue et al. 2008; Araç et al. 2006).
III.4 Synaptotagmin-‐1 promotes membrane fusion by inducing curvature
Finally, it was shown that Synaptotagmin-1 was able to bend membranes in the presence of calcium (Martens et al. 2007; Hui et al. 2009) and that this bending activity is mostly driven by the C2B domain (Hui et al. 2009). The positive membrane curvature induced by the C2B domain was able to stimulate liposomes fusion in concert with the SNARE proteins (Hui et al.
2009). Synaptotagmin-1 is therefore thought to function at least partially by performing bending work on the target membrane, providing a highly curved membrane to ease the high energy barrier associated with membrane fusion.
A" B" C"
15
All these models are probably ideal views that all have some truth. They are not necessarily incompatible and a mix of all or part of these models can be envisioned. However, it is clear that Synaptotagmin-1 interactions with membranes play a large role, and that insertion of the calcium binding loops of Synaptotamgin-1 in the membrane is the power stroke that triggers fusion.
IV. Previous measurements on Synaptotagmin-‐1 membrane interactions
Different types of approaches have been used to assess Synaptotagmin-1 binding to membranes, ranging from biochemical “bulk” assays to single molecule force spectroscopy.
Vesicle sedimentation assays have been traditionally used to measure the binding of soluble Synaptotagmin-1 (C2AB, without the transmembrane domain, with just the C2A and C2B domains) binding to vesicles. They estimate the affinity of protein-membrane binding at equilibrium by measuring the fraction of protein that binds to the vesicles as a function of the lipid concentration (Kuo et al. 2009).
Stopped-flow fluorescence resonance energy transfer (FRET), has also been used to study the binding of soluble Synaptotagmin-1 to vesicle through the monitoring of a FRET signal between the tryptophane residue of the protein and fluorescent lipids incorporated in the vesicle. Kinetics of binding (kon, koff), hence an affinity constant (Kd) can be calculated.
(Bai et al. 2004; Pérez-Lara et al. 2016).
An Atomic Force Measurement (AFM) (Takahashi et al. 2010) attached Synaptotagmin-1 to an AFM tip and performed adhesion force measurements on PS/PC membrane, but no binding energetics or distance-dependent Synaptotagmin-1-membrane interactions were reported.
Recently, the energetics of single-molecule C2AB interactions were measured with optical tweezers in the presence of calcium (Ma et al. 2017), but reported a lack of binding in the absence of calcium. Therefore, there remains a lack of direct probes of the long-range and
16
short-range behaviors and energetics of Synaptotagmin-1 binding at anionic membranes, especially with Synaptotagmin-1 embedded in a membrane, as occurs under physiological conditions. This is what we will do in this work.
V. Aims of this PhD
The methods presented in the previous section do not capture Synaptotagmin-1 binding to a membrane in the ideal geometrical and physiological conditions. Specifically, Synaptotagmin-1 interactions with membranes are important to probe in the physiological context of a Synaptotagmin-1 loaded vesicle approaching a target membrane. As the situation is too complex in vivo to truly isolate the Synaptotagmin-1 interactions, we will mimic this process in vitro.
Therefore, here we aim to provide unique information on Synaptotagmin-1 interactions with membranes. The main technique that we will use is a Surface Force Apparatus (SFA), which allows the direct measurement of the interaction energy between two surfaces as a function of their separation distance. The two surfaces are functionalized with bilayers, and Synaptotagmin-1 attached to one of them, mimicking a synaptic vesicle slowly approaching the plasma membrane of the neuron. Importantly, these measurements provide a membrane loaded with a physiologically-relevant area density of Synaptotagmin-1 approaching a target membrane that includes the primary lipids known to interact with Synaptotagmin-1. We obtain direct molecular level insights into the behaviors of Synapotagmin-1 during confinement, docking, and unbinding at an anionic membrane surface, including binding energetics, and molecular rearrangements. These insights will help reveal the still-mysterious role of Synaptotagmin-1 in synaptic transmission.
Chapter II will present the methods, focusing on the SFA principle and how it can be applied to the study of Synaptotagmin-1 interaction with membranes, including the deposition of lipid bilayers on the surfaces and the purification of a construct of Synaptotagmin-1 adapted to our study.
17
Chapter III will present a first set of SFA experiments performed on the wild type Synaptotagmin-1 to measure its energy of binding to anionic membrane. We will give direct measurements of this energy obtained in the absence or presence of divalent ions (magnesium and calcium) and propose a mechanism for possible configurations of Synaptotagmin-1 confined between the two membranes.
In chapter IV, we will repeat the SFA experiments of chapter III on multiple versions of Synaptotagmin-1 carrying mutations on specific membrane interacting sites, notably the calcium binding sites, in order to measure the impact of these mutations on the energetics and configuration of the protein between the two membranes.
18
19
Chapter II: The Surface Force Apparatus:
general principles and application to the measurement of Synaptotagmin-‐1 interactions with membranes
In order to have a new insight on the properties of Synaptotagmin-1, we propose to probe its interactions with membranes with a new approach: the use a Surface Force Apparatus (SFA).
This device was originally developed in the 1970s by Jacob Israelachvili, David Tabor, and coworkers (Israelachvili 1973; Israelachvili & Adams 1978; Israelachvili & Tabor 1972). This technique allows the measurement of the force between two surfaces as afunction of their separation distance. Its unique precision led to a considerable improvement of the comprehension of different fundamental types of interactions between extended surfaces (electrostatic, Van der Waals, solvation or steric forces…).
In this chapter, we will first describe the basic principles of the SFA (part I). Then, we will detail how we adapted this technique to the measurement of the interaction of Synaptotagmin- 1 to lipid bilayers. It required preparing supported bilayers (part II) that were tested on preliminary SFA force runs (III), and the purification of an appropriate construct of Synaptotagmin-1 (part IV).
20
I. The Surface Force Apparatus set up
I.1 General presentation of the SFA
The SFA used in the study in a reproduction of the “Mark II” device developed in 1978 by JacobIsraelachvili (Israelachvili & Adams 1978). This device allows the measurement of the force between two molecularly flat surfaces as a function of their separation distance, in air of in liquid. The force and distance are measured with an accuracy of 10-7 N and 0.1 nm respectively. A schematic representation of the apparatus is shown in figure II.1. We will describe the main elements of an SFA experiment: the preparation of surfaces, the surface displacement system and finally the force and distance measurement principles.
Figure II.1: Schematic representation of the Surface Force Apparatus (SFA) used in our study. Two silver coated mica surfaces are deposited on two lenses disposed in a cross cylinder geometry. White light reflexion between the surfaces creates a pattern of interference that gives a very accurate measurement of the distance between the two surfaces. Simultaneously the force is measured by the deflection of a cantilever spring. The overall set up gives the measurement of the free interaction energy E between the surfaces in function of their separation distance D. Extracted from (Perez et al. 2008)
The mica sheets (typical area of 8 9 8 mm
2) are glued with a thermosetting epoxy onto two cylindrical lenses, which are then arranged in a crossed-cylinder geometry. This geometry (equivalent to a sphere-plane geometry) makes the alignment easier, and enables testing many different surface regions. According to the Derjaguin approximation,
4the force, F(D), between these two curved surfaces is proportional to the interaction free energy per unit area, E(D), between two equivalent planar surfaces if the distance is much smaller than the radius of curvature, R, of the surfaces, and if the interaction energy decreases sufficiently rapidly with distance (at least in 1/D
2):
F ð D Þ =R ¼ 2 p E ð D Þ ð 3 Þ SFA measurements thus provide directly the inter- action free energy per unit area between two surfaces as a function of their separation distance.
The SFA remains a challenging technique. First of all, it is not easy to cleave mica and obtain a few square centimeters of thin sheets free of steps, of dust parti- cles, and of mica flakes. Then, upon gluing the mica onto the cylindrical lenses and mounting the lenses into the apparatus, care has to be taken to avoid dust particles or any other contamination to reach the surfaces. The liquids and the chemicals used during the experiment have to be ultra-pure in order to reduce sources of contamination. In addition, the SFA has to be installed on a vibration-free table to avoid noise in the force measurements, and in a thermostated room to reduce thermal drifts.
Mica surfaces can be coated with various mole- cules of interest (lipids, polymers, proteins), which makes it possible to study the complete interaction profile of these molecules using the SFA, and to identify the nature of the forces involved. From the mid-eighties on, several SFA studies of lipid bilayer interactions have been reported which aimed at characterizing the fundamental forces between mem- branes
9: van der Waals forces, double-layer forces, hydration forces, and undulation forces. Once the technique was routinely applicable to lipid bilayers, it was fairly straightforward to use functionalized lipid bilayers whose headgroups would carry the function to be studied,
19,25–27or to follow any structure modification of two interacting lipid lay- ers.
20When applied to interacting proteins (whose size is typically of a few nanometers), the angstrom resolution of the technique can provide the molecular detail of their assembly, including conformational change and passage through any intermediate bind- ing state.
15,22Cantilever spring Helical spring Leaf
spring
White light Prism
Spectrometer
Interference fringes
F(D)/R = 2πE(D)
Opposing mica surfaces
Cantilever spring Helical spring Leaf
spring
White light Prism
Spectrometer
Interference fringes
F(D)/R = 2πE(D)
Opposing mica surfaces
FIGURE 1. The Surface Force Apparatus (SFA).
Stiff cantilever Leaf spring
Flexible helical spring
±1µm
±1nm.
Stiffness ratio: 1000
FIGURE 2. The differential spring system.
PEREZet al.
242
21
I.2 Mica surfaces preparation
The surfaces from which we want to study the interaction profile are deposited on a thin sheet of mica. Mica is generally used because of the possibility to obtain molecularly flat surfaces of only a few micrometers thick and ~cm2 in area. This is done by successive cleavage of a thick mica sheet (about 1 mm) into thinner sheets until the desired thickness is reached. This step is critical because due to its high surface energy, the mica is easily contaminated, and any contamination by a particle would cause the failure of the subsequent experiment. Mica cleaving is done inside a laminar flow hood using a clean needle. The achievement of an appropriate thickness (~2 to 4 microns) is indicated when the reflection of light on the mica sheet shows colorful interferences. An area of uniform color is the signature of a constant thickness. We then cut squares of about 1 cm2 in these areas using a heated platinum wire.
These small mica pieces are deposited on a large and clean mica sheet on which they spontaneously adhere due to the Van der Waals forces, protecting them from further contamination. This large sheet is then back-silvered with ~55 nm of silver in an evaporator.
At the beginning of each experiment, the silvered mica surfaces are glued on two convex cylindrical lenses, the silver side facing the lens, which are then set up in the SFA chamber in a cross-cylinder geometry.
I.3 Surfaces displacement system
One of the two lenses is fixed on the upper side of the chamber. Its position remains constant throughout the experiment. The other lens is held by a bending spring used to measure the force. The apparatus contains a translation device that moves the lower lens toward the upper one employing two systems of distance control with increasing sensitivity. A micrometric translation or “coarse level” allows for a rapid positioning of the two surfaces with an accuracy of about 1 µm, typically at the beginning and at the end of the experiment. A finer level of displacement, with accuracy of 1 nm, is achieved by a differential spring system (figure II.2). It consists of a helical spring pressing against a cantilever spring one thousand times stiffer. When the helical spring is compressed by 1 µm, the cantilever bends 1 nm, moving the lower lens by the same distance.
22
Figure II.2: Differential spring system. Extracted from (Perez et al. 2008)
I.4 Distance measurement principle
The system composed by the two silver-coated mica surfaces forms a Fabry-Perot interferometer. When a white light is sent perpendicularly to the two surfaces, the various wavelengths are reflected and transmitted across the different interfaces. The silver coating on the mica surface is tuned so that only 1% of the incident light is transmitted, the remaining 99% being reflected. Only some of the wavelengths from the incident white light lead to constructive interferences. The transmitted light from the interferometer is sent to a spectrometer where it is split in several fringes, each fringe corresponding to a particular wavelength. These fringes are called Fringes of Equal Chromatic Order (FECO). In the spectrometer, they consist of lines for a planar interferometer, whereas they are curved in a curved interferometer (figure II.3). The wavelengths are characterized by their order of interference n. In an SFA experiment, the interference pattern used as a reference is the one obtained when the two surfaces are in contact (D=0) (figure II.4, top), for which we record the position of the fringes λ!!. When the surfaces are separated by a distance D, each fringe λn is shifted towards the longer wavelengths by Δλn compared with its original position (figure II.4, bottom).
This shift in the wavelength Δλn is linked to the separation distance D with the relation (Israelachvili 1973):
23 tan (2π
λ! µμD) = 2!!"#$! sin !!!!!!!!/!!
!!/!!!!! 𝜋 (1+(!!"#$! )!)cos (!!!!!!!!/!!
!!/!!!!! 𝜋)±(!!"#$! )!−1)
where
λn is the wavelength of the nth ordered fringe at distance D,
λ!! and λ!!!! are the wavelengths of the the n and n-1 ordered fringes respectively when the
mica interfaces are in contact
µmica is the refractive index of mica (=1.60) and
µ is the refractive index of the media sandwiched between the two mica sheets.
The + sign refers odd order fringes (n odd) and the - sign refers to even order fringes (n even).
The interference pattern can be observed either directly by eye at the exit of the spectrometer or more precisely with a CCD camera recording the exact position of the fringes of the whole spectrum.
Figure II.3: FECO of mica surfaces in contact (top) or separated by a distance D=10 nm (bottom). Each fringe appears as a doublet with a β- and a γ- component due to the birefringence of the mica.
D"="10"nm""
D"="0"nm""
odd" even"
24
I.5 Force measurement principle
The measurement of the force is based on the bending of the spring holding the lower surface.
During the measurement, the lower surface is moved toward the upper one at a slow, quasi- static velocity (typically 1-5 nm/s). Calibration is done at large distances where there is no interaction force by measuring the distance as a function of motor displacement. In that regime (no interaction force), the displacement of the mobile surface exactly corresponds to the displacement imposed by the motor. When attractive forces are present, the two surfaces move faster than the calibration. Conversely, when repulsive forces are present, the surfaces move slower than the calibration. The exact force can be calculated simply by applying Hook’s law:
F= k (Δx)
where k is the spring constant, and Δx is the actual displacement relative to the expected displacement. An illustration is shown in figure II.4.
The precision on the measurement of the force depends on the stiffness value of the spring:
for strong interactions, a spring of high stiffness is required whereas for the detection of weaker forces the use of a spring with lower stiffness is more adapted. The same spring was used in all our experiments, with a stiffness of 109 N/m. We estimate the precision on the force measurement in the range of 100 nN.
Figure II.4: Force measurement principle in the SFA. When a force appears between two surfaces (in this example, a repulsive force), the displacement imposed by the translational motor on the cantilever (dimposed) is different from the real displacement of the surfaces (dreal). The resulting force is given by F = k(dimposed-dreal).
dimposed
Cantilever spring of stiffness k
dreal
