# Quantum dot-based FRET nanosensors for talin-membrane assembly and mechanosensing

Audrey Ntadambanya, Julien Pernier, Violaine David, Kimihiro Susumu, Igor Medintz, Mayeul Collot, Andrey S. Klymchenko, Niko Hildebrandt, Isabelle Potier, Christophe Le Clainche, et al.

► To cite this version:

Audrey Ntadambanya, Julien Pernier, Violaine David, Kimihiro Susumu, Igor Medintz, et al. Quan-tum dot-based FRET nanosensors for talin-membrane assembly and mechanosensing. Angewandte Chemie International Edition, 2024, 63, pp.e202409852. 10.1002/anie.202409852. hal-04671321

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# Quantum Dot-Based FRET Nanosensors for Talin-Membrane Assembly and Mechanosensing

Audrey Ntadambanya, Julien Pernier, Violaine David, Kimihiro Susumu, Igor L. Medintz, Mayeul Collot, Andrey Klymchenko, Niko Hildebrandt, Isabelle Le Potier, Christophe Le Clainche, and Marcelina Cardoso Dos Santos*

**Abstract:** Understanding the mechanisms of assembly and disassembly of macromolecular structures in cells relies on solving biomolecular interactions. However, those interactions often remain unclear because tools to track molecular dynamics are not sufficiently resolved in time or space. In this study, we present a straightforward method for resolving inter- and intra-molecular interactions in cell adhesive machinery, using quantum dot (QD) based Förster resonance energy transfer (FRET) nanosensors. Using a mechanosensitive protein, talin, one of the major components of focal adhesions, we are investigating the mechanosensing ability of proteins to sense and respond to mechanical stimuli. First, we quantified the distances separating talin and a giant unilamellar vesicle membrane for three talin variants. These variants differ in molecular length. Second, we investigated the mechanosensing capabilities of talin, i.e., its conformational changes due to mechanical stretching initiated by cytoskeleton contraction. Our results suggest that in early focal adhesion, talin undergoes stretching, corresponding to a decrease in the talin-membrane distance of 2.5 nm. We demonstrate that QD-FRET nanosensors can be applied for the sensitive quantification of mechanosensing with a sub-nanometer accuracy.

## Introduction

Transmission of force between the cell and its environment is essential for the cell adhesion and migration required for tissue building and repair. Transmission of force across the cell plasma membrane is facilitated by focal adhesions (FAs), which are highly organised, self-assembled adhesive complexes.[1,2] FAs form where force transfer between the cytoskeleton and the environment is required. Some of FA compounds, including talin, are capable of sensing cytoskeletal tension.[3,4] Talin is a large (270 kDa) intracellular protein consisting of a head domain ($F_0-F_3$), a flexible neck, and a tail of 13-rod subdomains ($R_1-R_{13}$). Its head domain binds to the cell plasma membrane. On the one hand, it interacts with inositol phosphate groups[5,6] of phosphatidylinositol 4,5-bisphosphate (PIP$_2$) lipids at the plasma membrane, releasing talin autoinhibition.[7,8] Interestingly, it was demonstrated that $F_3$ interacts with $R_9$ and $F_2$ with $R_{12}$, resulting in an inactive snake-like biting its own tail conformation.[6,9] On the other hand, the transmembrane adhesion receptors integrins are also known to disrupt the autoinhibition interaction in cells.[10] Talin's head activates integrins and enhances cell attachment to the extracellular matrix.[11] The rod subdomains in talin's tail bind actin which polymerizes in filaments. Myosin II, the molecular motor, grafts filaments and applies tension by contracting them.[12] Consequently, talin's multiple helical bundle rod subdomains unfold, leading to the revelation of new protein binding sites,[13] allowing maturation of FAs.[4] Therefore,

[*] A. Ntadambanya, V. David, C. Le Clainche, M. Cardoso Dos Santos Université Paris-Saclay, CEA, CNRS, Institute for Integrative Biology of the Cell (I2BC), Gif-sur-Yvette, France
E-mail: marcelina.cardoso-dos-santos@i2bc.paris-saclay.fr

J. Pernier
Gustave Roussy Institute, Inserm U1279, Université Paris-Saclay,
Villejuif, France

K. Susumu, I. L. Medintz
Center for Bio/Molecular Science and Engineering U.S. Naval
Research Laboratory
Washington, USA

M. Collot, A. Klymchenko
Laboratoire de Bioimagerie et Pathologie, CNRS UMR 7021
Université de Strasbourg,
Strasbourg, France

N. Hildebrandt
Department of Engineering Physics, McMaster University,
Hamilton, ON L8S4L7, Canada

I. Le Potier
Centre de nanosciences et nanotechnologies (C2N), CNRS
UMR9001, Université Paris-Saclay,
Palaiseau, France

© 2024 The Authors. Angewandte Chemie International Edition published by Wiley-VCH GmbH. This is an open access article under the terms of the Creative Commons Attribution Non-Commercial NoDerivs License, which permits use and distribution in any medium, provided the original work is properly cited, the use is non-commercial and no modifications or adaptations are made.
talin can interact with numerous other FA components and form macromolecular assemblies.

The mechanism of talin unfolding, due to mechanosens-
ing, is not fully understood. For example, the orderly
unfolding of talin as a result of actin-myosin stretching has
never been revealed. Owing to the large size of talin,
resolving its unstretched conformation via crystallography
has not been accomplished either. Dedden et al. observed
that talin unfolds from a 15 nm globular arrangement to a
60 nm string-like structure via cryoelectron microscopy.[9]
Using magnetic tweezers, it was possible to artificially unfold
talin into a stretched conformation of up to 250 nm.[14]
However, that system did not account for the talin-
membrane interaction. Super-resolution microscopy on fixed
cells reported talin elongations of 180 nm[13] or even up to
350 nm.[15] Talin unfolding due to mechanosensing is there-
fore a topic of debate. Those examples show that probes
with better spatial and temporal resolution for tracking
molecular assemblies are required to understand how cells
sense the force of the cytoskeleton. However, various
mechanobioimaging techniques are already employed to
measure the tensile forces exerted on cells. For example,
cells are studied on different geometries (micropillars) or on
stiff or soft substrates using Traction Force Microscopy.[16,17]
The magnitude of the force is determined by the deforma-
tion of the substrate. Alternatively, the tension may be
quantified by aspiration through micropipettes or by meas-
uring shear forces using microfluidic techniques. Conven-
tional tension probes based on deoxyribonucleic acid
(DNA) are also in use. When a tensile force is applied to the
probe, a single-stranded DNA molecule undergoes unwind-
ing, thereby allowing the hybridization of a complementary,
dye-stained DNA strand.[18] To improve the comprehension
of cell mechanosensing, intermolecular FRET nanosensors
have been developed. FRET is a non-radiative energy
transfer that occurs between an excited state donor and a
ground state acceptor. It is widely applied for biosensing
and analysis of biomolecular binding.[19,20] Fluorescent
protein (FP) based FRET biosensors have already demon-
strated cytoskeleton-dependent stretching of talin.[21,22] A
pair of FPs is separated by a short spider silk peptide
between two structural regions: the head and the tail. These
force sensors have confirmed that talin is mechanosensitive
and measured forces around 1–20 pN. Additionally, FRET
pairs can be separated by a DNA linker. The Salaita group
engineered force-calibrated DNA-based FRET sensors to
report integrin tension in focal adhesion. These sensors
measure forces on a molecular scale (5–50 pN)[23,24] with
nanoresolution of ~25 nm.[18] However, the use of FP/dye-
based sensors has limitations. It may require delicate bleed-
through corrections, parallelization (one by one sensor), or
multi-read out (spectral coupled with lifetime) and be prone
to photobleaching.[19,20,25] Furthermore, the FRET pair must
be inserted in a restricted distance region to sense
cytoskeleton tension. Typically, donor and acceptor FPs are
separated by a short linker between two structural regions[21]
or delimit protein ends.[15,22] Although highly sophisticated,
these biosensors were unable to reveal the underlying
process of talin stretching. Certain studies have extracted

FA protein conformations,[26] but FRET with conventional
fluorophores has limited the distance range to circa 2 to
9 nm.[27] When using QD-based FRET nanosensors, this
distance range can be doubled and molecular distances can
be quantified with a spatial resolution far below the optical
diffraction limit.[28–31] The spectrally broad absorption, large
Stokes shift, and narrow emission of QDs are ideal for
efficiently removing background signals that can be very
problematic for FP-FRET biosensors. It is almost impossible
to excite only the FP FRET donor. Acceptor background
caused by direct light excitation, must be subtracted via
control experiments of using the acceptor FP alone. Where-
as FRET ratio (emission intensities of donor and acceptor)
measurements can alleviate the problem, the FRET ratio
does not readily provide quantitative distance information.
In contrast, QD-FRET nanosensors only need to measure
the photoluminescence intensity of the QD donor alone and
in the presence of the acceptor to directly assess donor-
acceptor distances.

In this study, we demonstrated that QD-FRET nano-
sensors can be applied for the quantification of biomolecular
interactions of FA proteins such as talin. We were able to
measure the separation distances between Talin and a giant
unilamellar vesicle (GUV) membrane with sub-nanometer
precision. This allowed us to deduce how larger or smaller
talin variants associate with the membrane. In addition, our
mechanosensing study showed that actin-myosin contraction
produced shortening of the distance between the GUV
membrane and talin tail end, suggesting an elongation of the
talin parallel to the membrane. The in vitro model used in
the study mimicked ventral actin-myosin stress fibers, which
are located on the adhesion cell surface and contract
between two successive FAs.[32] Indeed, ventral stress fibers
play a fundamental role in cell retraction and shape changes
during cell migration.

Results and Discussion

Minimal Focal Adhesion Autoassembles on GUV Membrane

The biomimetic in vitro model used in our study recon-
stituted a minimal FA composed of talin-1, a mechanosens-
ing protein, as well as actin and myosin cytoskeleton
proteins. Figure 1A provides a detailed overview of the step-
by-step implementation of the biomimetic assay. The
interaction between talin and the lipid bilayer was driven by
PIP₂ lipids, which were embedded in the GUV membrane.
Indeed, this protein-membrane interaction is required for
talin to undergo a conformational change, exposing new
binding sites for transmembrane integrins.[7,33,34] In addition,
GUVs contained cholesterol to mimic the plasma membrane
of FA, which is characterized by a high steroid content.[35]
To investigate membrane associated biomolecular assembly,
three variants of recombinant talin were produced based on
the rod subdomain structure. Each variant had a portion of
the head domain responsible for PIP₂ binding (F₂-F₃
domains). However, the talin variants differed in their actin-
binding ability and tail length (i.e., the number of rod
<figure><img src="image_2.png" /><figcaption>**Figure 1.** Talin variants associate with the membrane via PIP~2~. A) Biomimetic assay A~1~) GUVs are immobilized on a biotin-streptavidin functionalized surface via biotinylated lipids, A~2~) Talin conjugated with QD binds to GUVs containing PIP~2~ (dark green) through the head domain (yellow). Stretchable rod subdomains are represented in turquoise and a dimerization domain DD in pink. A~3~) To assemble a FRET sensor, a MemBright640 (purple) is added to the GUV membrane. A~4~) Filamentous actin (dark blue) binds to the end of talin tail and talin stretching can be induced by myosin (black) contraction. B) Talin AB–Ta variant comprises four rod subdomains, one of which (R~13~) binds to actin. The majority of talins undergo dimerization (right) through DD, whereas some remain monomeric (left). C) ΔAB–Ta variant comprises eight rod subdomains but no R~13~ D) mini ΔAB–Ta variant comprises three rod subdomains and no actin binding subdomain R~13~. All variants bind PIP~2~ through a part of head domain (F~2~–F~3~).</figcaption></figure>

subdomains, see Figures 1B,C,D). The only one actin binding variant, actin binding talin (AB–Ta, Figures 1B), was short and contained four rod subdomains (R~1–R~2~–R~3–R~13), including R~13~ known for primarily binding actin. [36] Indeed, it has been demonstrated via magnetic tweezers that pulling on R~13~ results in a response from the subdomains opposite to it, namely R~1–R~2~–R~3. [14] AB–Ta carried a dimerization domain DD, which allowed talin to form dimers (Figures 1B, right). However, our results (vide infra) revealed that a proportion of AB–Ta remained in a monomeric state (Figures 1B, left). The second variant, delta actin binding talin (ΔAB–Ta), lacked R~13~ and was unable to interact with actin, impairing its mechanosensing ability (Figures 1C). As it included eight rod subdomains (R~1–R~8), this was the longest variant studied. The third variant (mini ΔAB–Ta) shown in Figures 1D, lacked the actin binding subdomain and was the shortest with only three rod subdomains (R~1–R~2–R~3). Thus, our biomimetic model, which mimicked the membrane recruitment of talin and the mechanical stretching by actin-myosin, was ideally suited for investigating the molecular mechanism of talin mechanosensing.

## QD-based FRET Nanosensors Act as Nanomolecular Ruler

To quantify the inter- and intramolecular distances of Talin variants interacting with the GUV membrane, we developed QD based FRET nanosensors. Talin variants were functionalized using QDs (Figures 1A~2~, which bind to terminal hexahistidine (His~6~) motif at talin's tail end. MemBright640 acceptor was inserted into the GUV membrane (Figures 1A~3~). The hydrophobic chains of the MemBright640 probe insert into the outer lipid layer, leaving the hydrophilic head containing the dye in contact with the surrounding GUV buffer. The coupling of orange-emitting QD with red-absorbing organic fluorophore, MemBright640, resulted in a high spectral overlap *J* (Fig S1A) and an efficient FRET sensing. Narrow QDs emissions, with no red-shifted tail, allowed to collect photoluminescence intensity, avoiding a bleed-through leakage to the “acceptor channel” (Fig S1B). A similar FRET pair (QD605-Cy5) has previously been employed in a single QD-based DNA sensor.[28] In this study, talin-QD variants bound to GUV membrane and donor photoluminescence intensity (*I*~D~) was measured. Next, MemBright640 was added to the GUVs to measure quenched photoluminescence intensity of the donor in the
presence of acceptor ($I_{DA}$). Thus, FRET efficiencies $\eta$ were calculated using this equation:

$$ \eta = 1 - \frac{I_{DA}}{I_{D}} \qquad (1) $$

Since there was no spectral bleed-through between the donor and acceptor emissions, no specific correction of the $I_{DA}$ was required. Therefore, knowing both $\eta$ and the Förster distance $R_0$ allowed super-resolution of intermolecular interactions by measuring distances $r$ in molecular assemblies as follows:

$$ r = \left( \left( \frac{1}{\eta} - 1 \right) R_0^6 \right)^{\frac{1}{6}} \qquad (2) $$

With $R_0$ calculated as follows:

$$ R_0 = \left( \frac{9(ln10)\kappa^2\Phi_D J}{128\pi^5 n^4 N_A} \right)^{\frac{1}{6}} \qquad (3) $$

where $\kappa^2$ is the orientation factor, $\Phi_D$ is the quantum yield of the donor, $J$ is the overlap integral between donor emission and acceptor absorption, $n$ is the refractive index of the medium and $N_A$ is the Avogadro number. A critical parameter for obtaining $R_0$ is $\kappa^2$. Since $\kappa^2$ depends on the transition dipole moments of the donor emission and acceptor absorption, an imprecise knowledge of the orientation of the dipole moments can lead to a significant uncertainty in the distance calculation.[37] However, QDs provide a stable and durable orientation factor due to their isotropic emission. Here, the measured emission anisotropies for talin-QD associated to the GUV membrane and for Mem-Bright640 were 0 and 0.02, respectively. This confirmed the value of $\kappa^2=2/3$ for spherical QDs.[38] Consequently, calculated

$R_0$ was 6.6 nm for the used orange QD-MemBright640 FRET pair. This allowed to sensitively deduce donor-acceptor distances from the plot of $\eta$ over $r$ (Fig S2A) in a distance range between circa 4 nm and 11 nm (strongest change of $\eta$ with $r$). To achieve optimal acceptor insertion into the GUV membrane, we evaluated the integration of MemBright640 using fluorescence spectroscopy (Fig S2B). MemBright640 affinity towards the GUV was high, with an apparent dissociation constant (Kd) of 275 nM, and the GUV membrane reached saturation around 800 nM. Therefore, MemBright640 was used at 400 nM to obtain a stable intensity signal. The number of acceptors per single donor (A) must also be considered for distance calculation:[39]

$$ \eta = \frac{A * R_0^6}{A * R_0^6 + r^6} \qquad (4) $$

Based on a theoretical prediction of the mini ΔAB–Ta structure (see AlphaFold in Supporting Information), we assumed that $r=6.4$ nm. The $\eta$ found at 400 nM Mem-Bright640 was 0.58 (Fig S2C) which resulted in $A \sim 1$. Thus, our experiments were designed to easily quantify molecular distances using a 1:1 donor-acceptor ratio.

### Intermolecular Interaction Measured between Talin Variants and GUV Membrane is in the 6–9 Nanometer Range

First, intermolecular distances were measured using fluorescence spectroscopy. Talin variants were conjugated with QD donors and then mixed with GUVs to allow membrane-protein assembly. To prevent an excess of unbound QDs, QD was conjugated with talin variants at a ratio of 1:2 (Fig S3). QDs were excited at 405 nm, and the resulting photoluminescence spectra (Figure 2A,B,C, orange curves) showed their maximum intensity at ~600 nm. The interaction between talin and the GUV membrane was

<figure><img src="image_3.png" /><figcaption>**Figure 2.** Talin-GUV membrane interaction quantified using fluorescence spectroscopy. Normalized photoluminescence spectra of A) mini ΔAB–Ta, B) ΔAB–Ta and C) AB–Ta QD conjugated (donor) are shown by the orange curves. Normalized photoluminescence spectra of those talin-QD variants quenched by MemBright640 acceptor are shown by the blue curves. Corresponding $\eta$ are shown in black boxes. $\lambda_{ex}=405$ nm. D) Diagram of corresponding $\eta$ for three talin variants. The error bars indicate the standard deviation of the mean FRET efficiency. The results shown were confirmed in $N=4$ (mini ΔAB–Ta and ΔAB–Ta) or $N=3$ (AB–Ta) independent experiments.</figcaption></figure>
demonstrated upon the addition of MemBright640 acceptor, as illustrated by the quenched photoluminescence intensity (Figure 2A,B,C, blue curves). In addition to donor quenching, the acceptor showed weak sensitization at ~650 nm. Using the integrated areas of the donor and donor-acceptor spectra to calculate $I_D$ and $I_{DA}$, respectively, $\eta$ was determined using Eq. 1. Corresponding $\eta$ values of 0.48, 0.36, and 0.53, were found for the three talin variants, mini $\Delta$AB–Ta, $\Delta$AB–Ta and AB–Ta, respectively (Figure 2D, Table 1). Eq. 2 was used to determine the distance between QD donor and MemBright640 acceptor. For mini $\Delta$AB–Ta, $\Delta$AB–Ta and AB–Ta, the calculated distances were 6.7, 7.3 and 6.5 nm, respectively (Table 1). Since QD was at the tail end of talin, these distances reflected intermolecular distances separating talin from the GUV membrane. However, spectroscopic experiments were performed at the population level. We were therefore interested in performing experiments that could represent cell biology research at the single cell level. Since imaging allows to perform measurements of individual GUVs, we reproduced our experiments using fluorescence microscopy.

Similarly to spectroscopy, GUVs were incubated with talin-QD variants. To enable identification of the same GUV in two-step imaging (acquisition of $I_D$ and $I_{DA}$), GUVs were immobilized on biotin-streptavidin functionalized surfaces using biotinylated lipids incorporated in the GUV membrane (Figure 1A). First, images were captured in the “donor channel”, where QD donors were excited at $\lambda$= 405 nm and their intensities collected at $\lambda$=607 nm. Mean $I_D$ values were extracted from the membrane region for each GUV (Figure 3A₁, B₁, C₁). After the addition of Mem-Bright640, images were captured in the same channel to obtain $I_{DA}$ (Figure 3A₂, B₂, C₂). The most pronounced difference in intensity, between $I_D$ and $I_{DA}$ images, corresponded to a high $\eta$ (e.g., AB–Ta on Figure 3C₁ and C₂) which means that the shortening of the intermolecular distance made the quenching of the donor more visible. Prior to the addition of MemBright640, the sample served as non-biosensing control (Figure 3A₃, B₃, C₃). To compare with conventional FRET imaging, images were also acquired in the “biosensing channel”. QD donors were excited at $\lambda$= 405 nm, and emission from the MemBright640 acceptor was

<table><caption>**Table 1:** Comparison of FRET efficiencies $\eta$ and distances $r$ obtained using spectroscopy and microscopy for mini $\Delta$AB–Ta, $\Delta$AB–Ta, AB–Ta and AB–Ta stretched by actin-myosin. Sub-population (sb) 1 and 2 are sub-populations of values found using microscopy. $\delta$ are standard deviations. N.A. = not applicable.</caption><thead><tr><th rowspan="2"></th><th colspan="3">Spectroscopy</th><th colspan="3">Microscopy</th></tr><tr><th>η</th><th>δ<sub>η</sub></th><th>r (nm)</th><th>δ<sub>r</sub></th><th>η</th><th>δ<sub>η</sub></th><th>r (nm)</th><th>δ<sub>r</sub></th></tr></thead><tbody><tr><td>mini ΔAB–Ta</td><td>0.48</td><td>0.04</td><td>6.7</td><td>0.2</td><td>0.38</td><td>0.13</td><td>7.3</td><td>0.8</td></tr><tr><td>ΔAB–Ta</td><td>0.36</td><td>0.05</td><td>7.3</td><td>0.2</td><td>0.17</td><td>0.08</td><td>8.8</td><td>0.9</td></tr><tr><td>AB–Ta</td><td>0.53</td><td>0.16</td><td>6.5</td><td>0.7</td><td>0.44</td><td>0.19</td><td>6.9</td><td>0.9</td></tr><tr><td>AB–Ta sp1</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>0.34</td><td>0.08</td><td>7.4</td><td>0.5</td></tr><tr><td>AB–Ta sp2</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>0.71</td><td>0.07</td><td>5.7</td><td>0.4</td></tr><tr><td>AB–Ta + actin-myosin</td><td>0.63</td><td>0.04</td><td>6</td><td>0.1</td><td>0.74</td><td>0.17</td><td>5.5</td><td>0.8</td></tr><tr><td>AB–Ta + actin-myosin sp1</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>0.54</td><td>0.13</td><td>6.4</td><td>0.6</td></tr><tr><td>AB–Ta + actin-myosin sp2</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>N.A.</td><td>0.85</td><td>0.03</td><td>4.9</td><td>0.2</td></tr></tbody></table>

<figure><img src="image_4.png" /><figcaption>**Figure 3.** Talin-membrane interaction quantified using single-GUV fluorescence microscopy. Representative fluorescence images of three talin variants: A) mini $\Delta$AB–Ta, B) $\Delta$AB–Ta, and C) AB–Ta. In the “donor channel” photoluminescence intensity of the QD donor before (A₁, B₁, C₁) and after addition of the acceptor (A₂, B₂, C₂). In the “biosensing channel”, there was no interaction detected when the donor was alone (A₃, B₃, C₃). Alternatively, acceptor emission occurred due to the sensitization by the donor (A₄, B₄, C₄). “Donor channel”: $\lambda_{ex}$=405 nm, $\lambda_{em}$=607/36 nm, “biosensing channel”: $\lambda_{ex}$=405 nm, $\lambda_{em}$=692/40 nm, color bars below images represent intensity level, scale bars = 5 µm. D) Violin plot of $\eta$ for three talin variants. Each spot represents a single GUV and orange bars represent mean values with $\eta$=0.38 for mini $\Delta$AB–Ta, $\eta$=0.17 for $\Delta$AB–Ta, and $\eta$=0.44 for AB–Ta. Turquoise bars represent mean values of sub-populations with $\eta_1$=0.34 and $\eta_2$=0.71. The number of analyzed GUVs is indicated below the plots and the results presented were confirmed in N=3–4 independent experiments. Significantly different data (p < 0.01) are indicated by 3 stars. Statistical significance was tested by Kruskal-Wallis test followed by Conover paired comparisons.</figcaption></figure>
collected at $\lambda$=692 nm (Figure 3A₄, B₄, C₄). Differences in intensity were also observed, but were not quantified due to a relatively weak fluorescence signal, as already found in the spectroscopy experiments (Figure 2). Thus, $\eta$ for each individual GUV, represented by data points in the violin plot (Figure 3D) was calculated using Eq 1. For mini $\Delta$AB–Ta and $\Delta$AB–Ta, $\eta$ was 0.38 and 0.17, respectively (Table 1), and the violin plots showed a single population of values. In contrast, for AB–Ta, we found two sets of values: $\eta_1$=0.34 and $\eta_2$=0.71 (Figure 3D, Table 1). Since talin contains a dimerization domain, those two sets suggested that monomers and dimers of AB–Ta are not located at the same distance from the membrane. Further, the C-terminal 32 amino acids long His₆-tag, for QD binding, might contribute to the spread of data points within populations, as shown by the violin plot. For $\Delta$AB–Ta and mini $\Delta$AB–Ta, their His₆-tags were much shorter (8 amino acids). Kruskal–Wallis test followed by Conover paired comparisons confirmed the significant differences between $\Delta$AB–Ta and AB–Ta or mini $\Delta$AB–Ta (Figure 3D). To solve the orientation of talin with respect to the membrane, structural predictions were generated using AlphaFold machine learning approach.[40] However, not all of predicted structures were relevant as they were mostly based on the auto-inhibited talin structure and did not account for the talin-membrane interaction (see Supporting Information and Fig S4 and S5). The predicted distance for the mini $\Delta$AB–Ta was found to be the most accurate, given that this variant contains only three rod sub-domains. The data obtained from experimentation (7.4 nm) and predicted by the model (6.4 nm) were found to be the most in agreement. Consequently, predicted value was employed in order to calculate the number of acceptors per donor (Eq. 4). The data for AB–Ta and $\Delta$AB–Ta are less consistent with the experimental results (SI). In summary, the imaging experiments provided intermolecular distances of 7.4±0.5 and 5.7±0.4 nm for two sub-populations of AB–Ta variant, as well as 7.3±0.8 and 8.8±0.9 nm, for mini $\Delta$AB–Ta and $\Delta$AB–Ta variants, respectively (Table 1). As for the spectroscopy experiments, the distance increased from mini $\Delta$AB–Ta to $\Delta$AB–Ta. Imaging on a single GUV revealed information hidden within an entire population obtained via spectroscopy. $\eta$ values obtained by spectroscopy were higher than those obtained using microscopy. This could be explained by differences in experimental conditions. We assume that some of the GUVs exploded after the acceptor addition and settled in the cuvette, resulting in a $I_{DA}$ decrease and thus, a biased $\eta$.

## Mechanosensing Causes Talin to Organize Parallel to the Membrane

Actin and myosin II, two major filament components, have been used to mimic the tensile force of the cytoskeleton. To measure the intramolecular changes in talin length, we used the AB–Ta variant. AB–Ta-QD was assembled with the GUV membranes. AB–Ta variant contained actin binding subdomain $R_{13}$. Upon addition of myosin, actin filaments

contracted inducing AB–Ta stretching. Corresponding $I_D$ was measured using both spectroscopy and microscopy (Fig S6A, B₁). The FRET nanosensor was completed with MemBright640 acceptor and the quenching of the QD donor $I_{DA}$ was recorded (Figure 4A, S6A, B₂). To demonstrate the nanosensor insertion, images were recorded in both the “acceptor” and “biosensing channels” (Figure 4B, C). In the “actin channel”, actin bound to talin and polymerized along the GUV (white arrows, Figure 4D₂, D₃) which could be distinguished from a layer of filaments forming a network around the GUV. 3D images were reconstructed from z-stacks, and their top and lateral views are shown in Figures 4A₂, B₂, C₂, D₂ and 4A₃, B₃, C₃, D₃ respectively. Spectroscopy as well as microscopy experiments revealed a slight increase in a mean $\eta$ value (Figures 4E, F) between unstretched (left) and stretched (right) AB–Ta. As above, spectroscopy showed a net difference in distances, while microscopy revealed two distinct sub-populations. Thanks to spectroscopy, we were able to measure a $\eta$ displacement from 0.53 to 0.63, which corresponds to AB–Ta tail end approaching the membrane from 6.5 to 6 nm (Table 1). Measurements using microscopy were more precise with values of $\eta_1$=0.34 and $\eta_2$=0.71 for unstretched and $\eta_1$=0.54 and $\eta_2$=0.85 for stretched AB–Ta. This suggested that when talin is unstretched, some tail ends are located at 7.4±0.5 and others at 5.7±0.4 nm (Table 1). In contrast, when actin-myosin exerted tension on AB–Ta variant, tail ends relocated at 6.4±0.6 nm and 4.9±0.2 nm. Mean $\eta$ values (Figures 4F), as well $\eta_1$ and $\eta_2$ of sub-populations comparing stretched and unstretched conditions were significantly different. According to Gaussian distribution for QD: AB–Ta conjugated at ratio 1:2, the majority of AB–Ta variants were dimers, some remained monomers. This suggests that AB–Ta dimers undergo actin-myosin stretching, corresponding to a decrease in the distance between the tail end of talin and the membrane of 2.5 nm, from 7.4 (bottom left of the violin plot in Figures 4E) to 4.9 nm (top right of the same Fig). This decreased distance indicates a significant elongation of talin parallel to the membrane. In this scenario, actin binds to $R_{13}$ and myosin contracts the polymerized actin filaments along the membrane (Figures 5B₁), visible in Figure 4D₂ and D₃ (white arrows). The less widespread distance sub-population may correspond to AB–Ta monomers, i.e., 5.7 (top left of the violin plot in Figures 4E) extended to 6.4 nm (bottom right) but not parallel to the GUV membrane.

## Conclusion

In conclusion, we have demonstrated that QD-based FRET nanosensors can be used to quantify the assembly of FA proteins on a synthetic membrane. The distances between proteins and the membrane surface could be measured with an accuracy of less than 1 nm. Distances obtained using microscopy were more effective than those by spectroscopy in identifying differences within the same population. The microscopy results suggest that AB–Ta dimers undergo actin-myosin stretching (Figures 5A₁, B₁), corresponding to
<figure><img src="image_5.png" /><figcaption>**Figure 4.** Quantification of Talin mechanosensing. A–D) Fluorescence microscopy of FRET nanosensor of AB–Ta variant stretched by actin-myosin in A) the “donor channel” (photoluminescence intensity of the QD, λex = 405 nm, λem = 607/36 nm), B) the “acceptor channel” (fluorescence of MemBright640, λex = 642 nm, λem = 692/40 nm), C) the “biosensing channel” (donor excited and sensitized acceptor emitting, λex = 405 nm, λem = 692/40 nm) and D) the “actin channel” (fluorescence of actin labeled by Atto488, λex = 488 nm, λem = 525/45 nm). White arrows indicate actin-myosin fibers on the GUVs. A₁–D₁) Z-slice from a stack with few GUVs in the imaged area. A₂–D₂) Top view of the deconvolved 3D image of upper right GUVs on A₁–D₁. A₃–D₃) Lateral view of the deconvolved 3D image of the same GUVs. Color bars represent intensity level, scale bars = 10 µm. E) Diagram of η calculated from spectroscopy experiments for AB–Ta + /- Actin-Myosin. The error bars indicate the standard deviation of the mean FRET efficiency. The results presented were confirmed in *N* = 3 independent experiments. F) Violin plot of η for AB–Ta + /- actin-myosin. Each spot represents a single GUV, orange bars present η mean values with η = 0.44 (left, unstretched AB–Ta variant) and η = 0.74 (right, stretched AB–Ta variant). Turquoise bars represent η mean values of sub-populations with η₁ = 0.34 and η₂ = 0.71 (left, unstretched AB–Ta variant) and η₁ = 0.54 and η₂ = 0.85 (right, stretched AB–Ta variant). The number of analyzed GUVs is indicated below the plots and the results presented were confirmed in *N* = 2–3 independent experiments. Significantly different data (p < 0.001) are indicated by 3 stars. Statistical significance was tested by Mann-Whitney Statistics.</figcaption></figure>

a decrease in the distance between the distal end of talin and the membrane of 2.5 nm (from 7.4 to 4.9 nm). A second minor subpopulation, which we consider to be the AB–Ta monomers, exhibited distance extensions ranging from 5.7 to 6.4 nm (Figures 5A₂, B₂).

We showed that a minimal set of proteins, including talin, actin and myosin is able to initiate a mechanosensing process even in absence of talin binding partners such as vinculin. Vinculin is necessary to build a 3D scaffold and regulate the mechanical response of the adhesion. Two different mechanisms for linking talin with the actin-myosin machinery were proposed.[36,37] The first one stipulates that actin binds to R₁₃, then actin-myosin contracts and stretches R₂–R₃ at the opposite end, allowing vinculin binding. The second mechanism proposes the release of inhibition of R₂–R₃ by the vinculin direct binding. This would allow actin to bind to talin independently of R₁₃ (on other actin binding sites). Therefore, our results support findings that mechanosensing is likely initiated by the actin-myosin stretching initiating at R₁₃ subdomain, as determined via AFM and magnetic tweezers.[14,42] This initial mechanosensing step causes talin to become linearized and move closer to the membrane (Figures 5B₁). Super-resolution microscopy was

used to obtain information on the molecular conformation and geometry of talin in mature adhesions, where it is tilted at a 15° angle from the membrane. This revealed that talin determines the nanoscale architecture of the FA, acting as a molecular ruler.[4] Therefore, we propose that in nascent adhesions, talin is first immobilized in PIP₂-enriched nano-domains (Figures 5A₁). Due to the nanometric diameter of the PIP₂ clusters of approximately 10 to 50 nm, they could not be resolved under our diffraction-limited imaging conditions.[43] Talin is then captured by actin filaments that polymerize at the R₁₃ subdomain. These actin filaments are stretched by myosin in the vicinity of the membrane, at ~5 nm (Figures 5B₁). Eventually, the binding of vinculin results in the tilting of talin. In conclusion, QD-based FRET nanosensors have yielded quantitative insights into talin stretching dynamics. Using these tools, we will be able to elucidate emerging models for talin assembly with various partners. Furthermore, we intend to combine distance measurements with the quantification of cytoskeletal tension. The calibration of FRET versus force should enable the use of sensors in vitro and in cells.
<figure><img src="image_6.png" /><figcaption>**Figure 5.** Model of early talin mechanosensing in nascent adhesions in the example of AB–Ta. A₁) Talin dimerizes through its dimerization domain DD. The interaction with PIP₂ immobilizes talin and releases its autoinhibition. A₂) Some talins may remain monomeric. B₁) Actin binds to R₁₃, polymerizes, and actin-myosin stretches talin parallel to the membrane to expose R₂–R₃ for vinculin binding. B₂) Actin binds to talin, and actin-myosin contraction results in increase in distance of talin end from the membrane.</figcaption></figure>

**Supporting Information**

The authors have cited additional references within the Supporting Information (Refs. [44–51]).

**Acknowledgements**

This work was partially funded by the Interdisciplinary Object “OI Bioprobe” from Université Paris Saclay. It has benefited from Imagerie-Gif core facility supported by l'Agence Nationale de la Recherche (ANR-10-INBS-04/ FranceBioImaging; ANR-11-IDEX-0003-02/ Saclay Plant Sciences). Authors thank the Integrative Bioinformatics platform (BIOI2) for free use of AlphaFold@I2BC. Authors thank also Bruno Faivre and Clémence Vigouroux for purification of Talin variants.

**Conflict of Interest**

The authors declare no conflict of interest.

**Data Availability Statement**

The data that support the findings of this study are available from the corresponding author upon reasonable request.

**Keywords:** biophysics · nanoparticles · biosensors · cell adhesion · mechanosensing

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Manuscript received: May 24, 2024

Accepted manuscript online: July 15, 2024

Version of record online: September 12, 2024