A Fyn biosensor reveals pulsatile, spatially localized kinase activity and signaling crosstalk in live mammalian cells

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Version of Record published: February 4, 2020 (This version)
Accepted: January 8, 2020
Received: July 26, 2019

1. Of interest
Guanidine production by plant homoarginine-6-hydroxylases

Dietmar Funck, Malte Sinn ... Jörg S Hartig

Research Article Apr 15, 2024
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Abstract

Cell behavior is controlled through spatio-temporally localized protein activity. Despite unique and often contradictory roles played by Src-family-kinases (SFKs) in regulating cell physiology, activity patterns of individual SFKs have remained elusive. Here, we report a biosensor for specifically visualizing active conformation of SFK-Fyn in live cells. We deployed combinatorial library screening to isolate a binding-protein (F29) targeting activated Fyn. Nuclear-magnetic-resonance (NMR) analysis provides the structural basis of F29 specificity for Fyn over homologous SFKs. Using F29, we engineered a sensitive, minimally-perturbing fluorescence-resonance-energy-transfer (FRET) biosensor (FynSensor) that reveals cellular Fyn activity to be spatially localized, pulsatile and sensitive to adhesion/integrin signaling. Strikingly, growth factor stimulation further enhanced Fyn activity in pre-activated intracellular zones. However, inhibition of focal-adhesion-kinase activity not only attenuates Fyn activity, but abolishes growth-factor modulation. FynSensor imaging uncovers spatially organized, sensitized signaling clusters, direct crosstalk between integrin and growth-factor-signaling, and clarifies how compartmentalized Src-kinase activity may drive cell fate.

eLife digest

Cells contain networks of signaling proteins that can respond to a variety of cues from the surrounding environment. Often the cell’s response to these cues is not just controlled by the level of protein, but by changing the activity of signaling proteins. For example, a signaling protein in humans and other mammals known as Fyn regulates a number of different processes, including when a cell grows, dies, or develops a specialist role.

Defects in the activity of Fyn are associated with several diseases in humans including cancer and Alzheimer’s disease. However, it remains unclear how Fyn contributes to these diseases, or how the protein is able to precisely coordinate responses to multiple different cues in healthy individuals. This is largely because there are no readily available tools that are able to specifically detect where and when this protein is active in cells.

Researchers often use fluorescent proteins called biosensors as tools to detect where specific proteins are located in living cells over time. Now, Mukherjee, Singh et al. have developed a new biosensor named FynSensor to monitor the active form of Fyn in mammalian cells.

Microscopy imaging of FynSensor in several different cell types showed that although Fyn was present everywhere, it was only active in certain areas. In these areas the protein switched between an active and inactive state, with clear ‘pulses’ of signaling activity lasting a couple of minutes in response to specific cues. These areas of high Fyn activity behaved like signaling hubs in which several different cues integrate together before Fyn triggers an appropriate cell response.

These results shed light on how Fyn is able to precisely control many different processes in cells. In the future, FynSensor could be used to rapidly screen for drug-like molecules to treat cancer, Alzheimer’s disease and other conditions linked with defects in Fyn activity. Furthermore, the FynSensor could be adapted to allow researchers to study other signaling proteins in humans and other animals.

Introduction

Regulation of cell fate and behavior is achieved through complex and interconnected signaling networks acting in concert (Cai et al., 2014; Devreotes et al., 2017; Ridley et al., 2003). For precise signaling, activities of key signaling proteins are tightly compartmentalized in the cell, both spatially and temporally (Depry et al., 2015; Gulyani et al., 2011; Komatsu et al., 2011). Especially important to cellular controls are signaling nodes that function downstream of multiple receptor classes and help in signal integration. Src family kinases (SFKs) are such key signaling nodes; activated by cell adhesion receptors, integrins, receptor tyrosine kinases (RTKs, including growth factor receptors) and Gprotein-coupled receptors (GPCRs) among others (Abram and Courtneidge, 2000; Giannone and Sheetz, 2006; Grande-García et al., 2007; Parsons and Parsons, 2004; Thomas and Brugge, 1997). Src-kinases critically influence cell fate; regulating cell shape, migration and adhesion, survival and growth, stemness and differentiation making them important therapeutic targets in multiple diseases (Chetty et al., 2015; Gujral et al., 2014; Kim et al., 2009; Lewis-Tuffin et al., 2015; Nygaard et al., 2014; Saad, 2009; Timpson et al., 2001; Zhang et al., 2013; Zhang et al., 2014). Despite their importance, intracellular activity patterns of individual Src family kinases with spatial and temporal precision are still unclear. Visualization of active kinases in cells is essential to understand how Src kinases integrate signals and regulate multiple, and sometimes opposing processes, with fidelity and precision.

Currently available fluorescent SFK sensors have severe limitations. Most available biosensors, especially genetically encoded ones, do not directly report the intracellular distribution of active kinases since they rely on detecting the phosphorylation of a ‘pseudosubstrate’ peptide (Liao et al., 2012; Ouyang et al., 2008; Ouyang et al., 2019; Seong et al., 2011; Wang et al., 2005). As a result, biosensor readout – extent of phosphorylation of ‘sensor’ peptides – can be affected by the activities of kinases as well as cellular phosphatases. Confounding readouts further, it has been reported that sensors can get trapped in an ‘ON’ state owing to strong intramolecular interactions between phosphorylated substrate peptides and their respective recognition motifs (Komatsu et al., 2011; Regot et al., 2014); and therefore are unable to report the turning OFF of kinase activity. Another major drawback of current sensors is that multiple kinases, especially closely related ones, can phosphorylate these substrate-based sensors in a promiscuous manner leading to a lack of specificity.

Study of SFK activity is especially complicated by the presence of multiple closely related Src family members, including three ubiquitously expressed kinases c-Src, c-Yes and Fyn. While there is some functional redundancy, individual SFKs also perform critical, exclusive roles in the cell (Zhang et al., 2014; Kuo et al., 2005; Lowe et al., 1993; Lowell and Soriano, 1996; Marchetti et al., 1998; Molina et al., 1992; Palacios-Moreno et al., 2015). Therefore, despite decades of study, it is not always clear which kinase(s) is activated in a given cellular context? In this regard, fluorescent biosensors that report the activation of individual Src kinases in live cells would be extremely valuable in clarifying kinase activity and function. However, such specific biosensors for individual Src kinases are still limited (Gulyani et al., 2011; Koudelková et al., 2019; Paster et al., 2009; Stirnweiss et al., 2013), with current sensors generally giving only a pan-SFK readout with limited spatial and temporal resolution.

Here, we present a fluorescent biosensor specific for the SFK-Fyn. Fyn, is ubiquitously expressed and regulates cell migration, epithelial to mesenchymal transition (EMT), cancer metastasis, immune-response, axonal guidance and patterning, and synaptic functions (Gujral et al., 2014; Du et al., 2016; Lewin et al., 2010; Meriane et al., 2004; Posadas et al., 2016; Salter and Kalia, 2004). Interestingly, among several SFKs implicated, recent evidence shows Fyn may specifically control EMT and metastatic progression (Gujral et al., 2014). Overall, Fyn is a critical player in multiple cell/tissue types, integrating signaling through multiple receptor classes (Palacios-Moreno et al., 2015; Martín-Ávila et al., 2016; Yadav and Denning, 2011) and controlling cell fate. However, there is currently no tool, including specific antibodies, to visualize Fyn activity in cells. As a consequence, there is no information on intracellular Fyn dynamics in live cells. A Fyn biosensor would bridge this gap and offer a method to address Fyn functional complexity as a key signaling node.

Generation of biosensors is often limited by a lack of naturally available binders that can detect active states of proteins in real-time. To address this, we have developed a strategy of using artificially engineered binders that recognize the active form of target proteins in live cells (Gulyani et al., 2011). Like other SFKs (Arold et al., 2001; Boggon and Eck, 2004; Kinoshita et al., 2006; Noble et al., 1993; Sicheri and Kuriyan, 1997), Fyn is maintained in an inactive or closed form through multiple intramolecular interactions (Thomas and Brugge, 1997). Activation of the kinase involves disruption of these interactions leading to an open conformation, with more exposed surfaces (Gulyani et al., 2011; Young et al., 2001). Taking advantage of this, we screened a combinatorial protein library to isolate a binding protein (F29) that specifically binds to a region of Fyn (SH3 domain) that is more exposed in the open conformation. For screening, we chose a library generated by mutagenizing the Sso7d protein - a highly stable protein derived from the hyperthermophilic archaeon Sulfolobus solfataricus. We have previously shown that Sso7d mutant-libraries can be used to isolate binding proteins for a wide range of targets (Gera et al., 2012; Gera et al., 2011; Gocha et al., 2017; Hussain et al., 2013). Also, since Sso7d has no known interactions with mammalian intracellular proteins, we reasoned that Sso7d would serve as an ideal ‘inert’ scaffold for generating an intracellular biosensor. Our analysis shows that F29 indeed binds specifically to Fyn, with little to no cross-reactivity with other SFKs. Nuclear magnetic resonance (NMR) spectroscopy analysis of F29 in complex with the target reveals the structural basis of the specificity of F29 binding to Fyn.

An efficient readout of the molecular recognition of the target by the binding protein is central to constructing an effective biosensor. Here, we used our new binder/F29 to construct a genetically encoded Fyn biosensor (FynSensor) based on fluorescence resonance energy transfer (FRET). FynSensor is robust, sensitive and faithfully reports Fyn activation and regulation. We also show how FynSensor expression in cells is minimally perturbing, with no measurable changes in either cellular morphodynamics or downstream signaling. Significantly, FynSensor enables direct visualization of active Fyn in live cells, revealing Fyn activation to be spatially compartmentalized, polarized and pulsatile. To the best of our knowledge, this is the first reported visualization of activated Fyn in live cells; notably, unavailability of suitable antibodies currently precludes even routine detection of active Fyn. Strikingly, FynSensor imaging shows the presence of spatially localized signaling clusters sensitive to integrin and growth factor signaling; and reveals how growth factor response in cells is influenced by integrin-dependent protein activity. More broadly, FynSensor shows how spatially compartmentalized activation of key signaling proteins may lead to efficient signal integration and precise control of cell physiology. Further, our results provide a framework for systematic development of intracellular biosensors for specific Src-family and other kinases in general.

Results

Screening yields binder specific for active Fyn

Specifically recognizing active Fyn is critical for constructing an intracellular sensor. Activation of SFKs is accompanied by large conformational changes; the active kinase adopts a more open conformation with significantly reduced intramolecular interactions (Thomas and Brugge, 1997). Specifically, prior evidence (Gulyani et al., 2011; Young et al., 2001) suggests that the SH3 domain in SFKs is more accessible in the active conformation relative to the closed, inactive state (Figure 1A). Therefore, a binding protein targeting the SH3 domain of Fyn can be expected to preferentially bind the active form of the kinase (Gulyani et al., 2011). Such an active state binder can then be engineered as a fluorescent biosensor for reporting intracellular Fyn activation (Figure 1A,B). Accordingly, we sought to generate a binding protein specific for Fyn SH3 domain. To this end, we screened a yeast surface display library of Sso7d mutants (library generated by randomizing ten Sso7d residues), using magnetic screening and fluorescence associated cell sorting (FACS), adapting methods described previously (Gera et al., 2011; Gera et al., 2013) (Figure 1—figure supplement 1A,B). Stringent negative selection steps were included to eliminate binders that exhibited cross-reactivity to the SH3 domains of seven other SFKs, specifically c-Src, c-Yes, Fgr, Blk, Lck, Lyn, and Hck. The pool of yeast cells obtained after magnetic selection were further subjected to multiple rounds of FACS to enrich Fyn SH3-binding yeast cells (Figure 1—figure supplement 1C). Single clones were then picked and assessed for specific binding to Fyn-SH3. One such clone (referred to as F29 hereafter) showed specific binding to Fyn-SH3 domain, with little or no cross-reactivity with SH3 domains from the other highly homologous SFKs mentioned above (Figure 1C and Figure 1—figure supplement 2). Sequence of F29 and its comparison to wild-type Sso7d is shown in Figure 1D.

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High-throughput screening yields a new protein binder for generating a specific biosensor for active Fyn.

(A) Rationale for a biosensor targeting active conformation of SFK, Fyn. A cartoon depicts how activation of Fyn kinase leads to a more open conformation and buried regions (including its SH3 domain) becoming more accessible for biosensor binding. Inactive (*left*) SFKs are held in a closed conformation through two key intramolecular interactions (for instance see *Cell*, 2001, 105, 115–12) that are lost in the open state (*right*). (B) Schematic diagram representing sensing of active and open state of kinase by an active state sensor. (C) High-throughput combinatorial library screening yields a protein binder specific for Fyn SH3 domain. Analysis of the yeast clone ‘F29’ using fluorescence activated cell sorting (FACS) shows F29 preferentially binds Fyn while showing little or no binding to SH3 domains of ubiquitously expressed SFKs, c-Src and Yes (1 µM each). Here, fluorescence signal due to biotin-labeled target SH3 (Streptavidin-PE) is plotted against signal due to yeast cells expressing the c-myc-tagged Sso7d scaffold (Alexa 633 labeled anti-c-myc). Detection of fluorescence signal due to Fyn-SH3 along with anti-c-myc fluorescence in a single yeast cell, indicates expression of full-length F29 fusion-protein and binding to labeled Fyn-SH3 target (also see Figure 1—figure supplement 2). (D) Comparison of amino acid sequences of new binder F29 with original Sso7d scaffold. 10 Sso7d residues randomized for library generation are shown in bold while corresponding new residues in binder F29 are shown in blue. (E) Binder F29 is able to recognize Fyn SH3 outside the context of yeast surface with recombinantly expressed and purified F29 showing saturable binding to Fyn SH3. Immunoblot showing pull-down of His-tagged Fyn or Src SH3 domains (75 nM to 1.25 µM) using GST-F29 fusion protein immobilized as bait on glutathione sepharose beads. His-tagged SH3 protein detected via anti-6X-His antibody (also see Figure 1—figure supplement 3). (F) Amount of bound Fyn SH3 plotted against total input Fyn SH3. The data from three independent binding experiments (n = 3) was plotted and fitted assuming 1:1 binding using a GraphPad Prism module. The apparent K_D values were calculated using Equation 1 (refer Materials and methods). Values are mean ± s.e.m.

Figure 1—source data 1 Analysis of F29 binding to Fyn-SH3.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig1-data1-v1.xlsx
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We further examined if F29 retained binding to Fyn-SH3 when removed from the context of the yeast cell surface. F29 fused with glutathione-S-transferase(GST-F29) was recombinantly expressed and purified, and immobilized on glutathione sepharose beads. Immobilized GST-F29 could pull down 6xHistidine (6xHis) -tagged Fyn-SH3, as detected by immunoblotting using an anti-6xHis antibody. In contrast, no signal was seen when the Fyn SH3 pulldown experiment was done with GST-saturated beads, or when the SH3 domain of Src was pulled down using GST-F29 beads (Figure 1—figure supplement 3, Figure 1E). Taken together these results confirm that F29 binds specifically to the Fyn-SH3. We further used pulldown experiments to estimate the binding affinity (K_D) of the interaction between F29 and Fyn-SH3. Briefly, immobilized GST-F29 was incubated with varying concentrations of Fyn-SH3, and the amount of bead-bound Fyn-SH3 was quantified. Upon fitting the data to a monovalent binding isotherm, the apparent K_D of the F29-Fyn-SH3 interaction was estimated as 60 ± 16 nM (Figure 1F, Figure 1—source data 1).

Molecular basis of specific Fyn recognition by F29

F29 specifically binds Fyn SH3 while showing little or no cross-reactivity to SH3 domains of other highly homologous SFKs. To investigate the molecular basis of the specificity of F29 binding to Fyn-SH3, we used NMR spectroscopy to determine the structure of F29 in complex with Fyn-SH3. For this, we first analyzed the structure of isolated F29 (Figure 2). The ¹⁵N-edited Heteronuclear Single Quantum Correlation (HSQC) spectrum of purified F29 (isolated from E. coli grown in ¹³C, ¹⁵N-labeled media) showed well-dispersed peaks, indicating proper folding of the protein despite extensive mutagenesis of the Sso7d scaffold (Figure 2—figure supplement 1A). Complete chemical shift assignments of the backbone and side chain atoms using a series of 2D and 3D NMR experiments were then used to calculate a structure by the CS-ROSSETTA algorithm (van der Schot et al., 2013). The structure of F29 superimposed well with Sso7d (Figure 2—figure supplement 2A) indicating a conservation of the Sso7d fold in the binder.

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Structure and binding interface of Fyn-SH3:F29 complex by NMR.

(A) The lowest energy structure from Haddock is provided, where Fyn-SH3 is colored green and F29 is colored in light blue. (B) The surface representation of A, where the surface of Fyn-SH3 and F29 are colored in green and light blue, respectively. (C) *Ribbon diagram of contact interface*: A view of the contact interface where the F29 surface is represented in light blue and Fyn-SH3 is shown in green ribbon. The interacting side chains of Fyn-SH3 are shown in yellow. A key interacting residue, Asp99, that forms a salt-bridge with F29 and has been subjected to mutagenesis is shown in red. (D) *Complementary view of interface*: Complementary view of the interface with Fyn-SH3 in green and F29 in blue (ribbon diagram) is shown. The interacting side chains of F29 residues are shown and identified here. They are colored in dark red if randomized and colored in gold otherwise. The non-randomized residues Arg25 and Phe32 interact with Fyn-SH3 through the backbone atoms and hence, their ribbons are colored in gold. Non-interacting regions of F29 far away from the interface are hidden for clarity. (E) Schematic showing Fyn-SH3:F29 amino acids involved in interfacial contacts. The hydrophobic and aromatic-aromatic contacts between specific residues are shown in black lines, the hydrogen bonds are shown in blue lines and salt-bridges are shown in red lines. Residues of F29 are colored in red if randomized and colored in black otherwise. Asp99 in Fyn-SH3 is colored in red (Also see Figure 2—figure supplement 3B).

To obtain the structure of Fyn-SH3:F29 complex, ¹⁵N-edited HSQC spectra of ¹⁵N-labeled free Fyn SH3 domain (residues 87 to 139 of h-Fyn) as well as in complex with F29 (unlabeled) were acquired. The differences between chemical shifts in the free and bound Fyn-SH3 provided the Chemical Shift Perturbations (CSPs) in Fyn-SH3 due to F29 binding (Figure 2—figure supplement 1B,D). High CSPs point to specific Fyn SH3 residues likely present at the binding interface in the complex with F29 (Figure 2—figure supplement 1D). Similarly, the CSPs in¹⁵N-labeled F29 upon binding Fyn-SH3 were also obtained (Figure 2—figure supplement 1A,C); these values help identify the F29 binding interface in the complex. A 3D ¹⁵N-edited Nuclear Overhauser Effect Spectroscopy (NOESY) experiment, performed on a complex of ¹⁵N, ²H-labeled Fyn-SH3 and unlabeled F29, led to the unambiguous assignment of ~7 intermolecular NOEs from the spectra. These NOEs helped identify through-space interfacial interactions between residues. Using the structure of Fyn-SH3 (pdb: 3UA6), the structure of F29 obtained above, and the CSPs as ambiguous restraints, and NOEs as unambiguous restraints, the structure of Fyn-SH3:F29 complex was solved by Haddock (de Vries et al., 2010; van Zundert et al., 2016).

After analysis, the most populated cluster had 200 structures (Supplementary file 1A) and the lowest energy structure and its surface representation are shown in Figure 2—figure supplement 2B and Figure 2A,B. The contact residues at the interface of the complex are shown in Figure 2C,D. At the binding interface several non-covalent interactions including hydrophobic-aromatic, hydrogen bonds and salt bridges can be observed (Supplementary file 1B). Eight of the ten randomized residues as well as five originals (SSo7d scaffold) amino acids of F29 show interactions with twelve amino acids of Fyn SH3 (Figure 2E). Analysis of the binding interface reveals that residues Phe22, Thr24, Tyr26, and Phe31, 32, 43 of F29 show multiple interactions with the Fyn SH3 domain, with SH3 residues Tyr137, Asn136, Tyr132, Trp119, Arg96 and Tyr91 featuring prominently in these interactions. Interestingly, Lys9 (an original Sso7d residue) and Arg33 (randomized residue) on the F29 binder showed exclusive salt bridge interactions with Asp118 and Asp99 of Fyn SH3 domain, respectively (Figure 2E).

SH3 domains naturally interact with proteins with poly-proline motifs (Li, 2005). To gain insight into how F29 binds and specifically recognizes Fyn SH3, we compared the F29-Fyn SH3 complex with structures of FYN-SH3 bound to poly-Proline peptides (pdb: 4EIK and pdb: 3UA7) (Figure 2—figure supplement 1E,F). The buried surface area of F29:Fyn SH3 complex is much larger (800 Å², ∆G = −7 kcal/mol) compared to that of the poly-Pro:Fyn SH3 complex (pdb: 3UA7, 330 Å², ∆G = −5 kcal/mol). The number of contacts is also considerably larger in the F29 (84 contacts) versus the poly-Proline (40 contacts). Moreover, the orientation of the poly-Prolines at the interface differs from the orientation of the F29 interfacial beta-sheet by ~30^o. Hence, although the F29 interface overlaps with the poly-Proline interface, it is significantly distinct in terms of contacts, surface area and orientation. Residues of Fyn-SH3 that are involved in binding F29 as well as Poly-Pro-1/2 polypeptides are shown in Figure 2—figure supplement 1G.

We then used site-directed mutagenesis to examine contributions of key interfacial residues to F29-Fyn SH3 binding. A salt bridge between the residue R33 of F29 and D99 of Fyn SH3 appears prominent. Indeed, the introduction of an R33A mutation in F29 resulted in reduced binding to Fyn-SH3 domain, as evidenced by pull-down analysis (Figure 2—figure supplement 3A, Figure 2—figure supplement 3—source data 1). These results confirm that R33 in F29 and D99 in Fyn-SH3 contribute substantially to the F29-Fyn-SH3 binding interaction. Interestingly, a Pro41Ala mutant of F29 also showed reduced binding to Fyn-SH3 (Figure 2—figure supplement 3A). Although the NMR analyses do not show Pro41 to form direct interfacial contacts with SH3, the reduced binding of this mutant may be attributed to perturbation of local structure. Proline41 forms a critical hinge preceding β3 strand (Figure 2D); mutation of the proline likely disrupts the structure the β-sheet that contains multiple key binding contacts with Fyn-SH3.

These structural analyses also offer insight into the molecular basis for the specificity of F29 binding to Fyn-SH3 over highly homologous SH3 domains of other SFKs. While the binding interface in the Fyn-SH3-F29 complex reveals multiple interactions (Figure 2D,E), the residue D99 in Fyn-SH3 that forms a salt bridge with R33 of F29 is unique to Fyn among the ubiquitously expressed SFKs as well as Lyn and Fgr (Figure 2—figure supplement 3B). Therefore, we hypothesized that D99 in Fyn-SH3 may contribute to the binding specificity of F29 for Fyn-SH3 vis-à-vis to at least some of the SFK SH3 domains. To test this hypothesis, we generated a mutant version of the SH3 domain from Src (Src-SH3) wherein the native threonine was replaced with aspartic acid (T99D). Strikingly, T99D Src-SH3, but not wild-type Src-SH3, showed detectable binding to F29 in pulldown assays (Figure 2—figure supplement 3C). These results suggest that D99 in Fyn-SH3 contributes to the binding specificity of F29 for Fyn over other SFKs, notably Src. Nevertheless, specificity of F29 for Fyn-SH3 over Yes-SH3 (containing E99) (Figure 1C,E) and Lyn/Fgr (containing D99, Figure 1—figure supplement 2), suggests that binding specificity results from a unique combination of multiple interactions at the binding interface that may require further investigation.

F29 enables construction of FynSensor, a FRET biosensor for active Fyn

F29 efficiently and specifically binds Fyn SH3 in vitro. We then examined if F29 is able to recognize full-length, cellular Fyn and if it preferentially binds the active form of Fyn. To test this, either WT Fyn kinase or the CA mutant (Takeuchi et al., 1993) (active and open conformation) was exogenously expressed in human embryonic kidney (HEK-293T) cells and the cell lysate treated with immobilized GST-F29. Fyn pulldown and immunoblotting showed that the CA mutant was more efficiently pulled down by F29 as compared to the WT Fyn kinase (Figure 3A, Figure 3—source data 1). On the contrary, GST-alone control beads showed no significant pulldown of either WT or CA Fyn. These data indicate that F29 preferentially binds active Fyn; the SH3 domain targeted by F29 is more accessible in the CA mutant compared to the WT Fyn kinase. Further, similar GST-pulldown and immunoblot analysis showed that immobilized F29 (GST-F29), but not GST-alone control, was able to bind and pull-down endogenous or native Fyn from cell lysates (Figure 3B). These results confirm efficient binding of F29 to cellular Fyn.

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Developing a FRET biosensor for sensing active Fyn in cells.

(A) F29 preferentially binds active Fyn kinase. (I) Immunoblots showing GST-F29 pulldowns of cellular Fyn after transfecting cells with either Fyn WT (wild type) or Fyn CA (constitutively active kinase). (II) Extent of pulldown with either CA or WT Fyn. The band intensity of pulled-down Fyn kinase from five such experiments (n=5) was quantified and normalized to expression level. Binder F29 is able to pull-down CA mutant of Fyn to a greater extent than WT. Values are mean ± s.d from five independent experiments (n=5). Data was compared using Student’s unpaired one-tailed t-test on GraphPad Prism. (B) F29 binds full-length, endogenous cellular Fyn kinase. Immunoblots, using anti-Fyn antibody, show pulldown of full-length endogenous Fyn from cell lysates by GST-F29 immobilized on glutathione beads (n = 3). GST-coated beads are shown as controls. Coomassie stained gels of the bait proteins, either GST-F29 or GST used for pulldown are also shown. (C) Schematic diagram and model for an intermolecular FRET biosensor for visualizing active Fyn in cells. The donor fluorophore (mCerulean) is inserted in between Fyn kinase unique (UD) and SH3 domains using flexible linker sequences to minimally disrupt kinase function (also *see* Figure 3—figure supplements 2 and 5). Newly developed Fyn active-state binder, F29, is fused with acceptor fluorophore (mVenus). In the cell, F29 binder (FRET-acceptor) would bind the exposed SH3 domain of active Fyn (FRET donor), resulting in an intermolecular FRET between donor and acceptor fluorophores (*bottom panel*). In contrast, F29 would be expected to show significantly lower binding and reduced or no FRET when Fyn is inactive (*upper panel*). (D) Sensing active Fyn in cells through binding-dependent FRET. Fluorescence spectral measurements from HEK-293T cells co-transfected with FRET-donor (WT or CA Fyn-mCerulean) alone OR along with a FRET-acceptor (either Fyn binder mVenus-F29 or the non-binding mutant, mVenus-F29P41A). Specifically, the normalized emission spectra of transiently transfected cells when excited at 435 nm (corresponding to donor mCerulean excitation) are shown. Spectra has been normalized relative to emission intensity at 475 nm (corresponding to mCerulean emission). Overall, the spectra show that FRET (‘sensitized’ acceptor emission at 525 nm) is greater for CA Fyn as compared to WT Fyn. No FRET signal is observed (sensitized 525 nm emission) with donor alone or non-binding mutant F29-P41A. *Left inset figure* shows that binding and increase in FRET efficiency is saturable. Using raw fluorescent intensity values the FRET efficiency was calculated as per the equation Fa/(Fa+Fd), where Fa is acceptor and Fd is donor raw fluorescence intensity, respectively. Increasing the amount of labeled binder (mVenus-F29/acceptor) in cells while maintaining constant Fyn (donor) levels causes an increase in FRET till saturation is reached (blue closed squares), indicating binding dependent FRET. From three independent experiments (n=3) , a nonlinear best fit was plotted (R² = 0.97) using GraphPad Prism (blue, dotted line). In contrast, acceptor-labeled non-binder control (F29 P41A mutant) shows little or no FRET (orange open symbol) showing FRET response is binding dependent and specific. *Right inset figure* shows a bar plot of FRET efficiency comparing WT and CA Fyn for a 1:2 donor:acceptor molar ratio. From three such independent experiments (n=3) , data was compared using Student’s unpaired one-tailed t-test on GraphPad Prism. Values are mean ± s.e.m. (E) Membrane tagging of F29 increases biosensor sensitivity and FRET efficiency (calculated as above). (I) The emission spectra of cells co-transfected with Fyn (WT) and, either mVenus-F29 or myr-mVenus-F29 at a 1:4 donor:acceptor molar ratio on 435 nm excitation are shown. Spectra for corresponding non-binding controls (mVenus-F29P41A with and without myristoyl tags) are also shown. (II) Box-and-Whisker plot showing observed FRET efficiency (calculated as above) in cells when myristoyl-tag is added to the FRET-acceptor (mVenus-F29) or its non-binding control mutant P41A. Here, Whiskers represents the lowest and highest observed values of FRET efficiency, while Box spans the interquartile region with the ends demarcating the lower and upper quartile values. The mid-line marks the median. Plus symbol (+) represents the mean FRET efficiency. Data from four such independent experiments (n=4) were analyzed using Student’s unpaired one-tailed t-test on GraphPad Prism (respective p values are indicated on panel).

Figure 3—source data 1 Analysis of F29 pulldown of Fyn and its constitutively active CA mutant.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig3-data1-v1.xlsx
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Figure 3—source data 2 Analysis of FynSensor FRET efficiency with changing the cellular levels of acceptor F29 and non-binding control.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig3-data2-v1.xlsx
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Figure 3—source data 3 Analysis of FynSensor FRET response towards WT Fyn and constitutively active (CA) Fyn.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig3-data3-v1.xlsx
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Figure 3—source data 4 Analysis of biosensor FRET efficiency observed with various labeled F29 and non-binding constructs.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig3-data4-v1.xlsx
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Taken together, our results so far show F29 to be a promising candidate for generating an intracellular biosensor for active Fyn. Accordingly, we conceptualized a biosensor design based on FRET wherein the binding of F29 to active Fyn would result in an increase in FRET between suitably placed donor and acceptor fluorophores (Figure 3C). To construct a genetically encoded biosensor, Fyn was labeled with a FRET donor fluorescent protein (mCerulean) while F29 was tagged with a FRET acceptor protein (mVenus). While F29 already shows high specificity to Fyn, our design further ensures that FRET signal can arise only from labeled F29 binding to labeled active Fyn. For generating the fluorescently labeled Fyn, we chose to introduce mCerulean (FRET donor) between the unique (UD) and the SH3 domains of full-length Fyn kinase (Paster et al., 2009; Stirnweiss et al., 2013) (Figure 3C). The choice of this insertion position is designed to increase the probability of efficient FRET when acceptor-labeled F29 binds the exposed SH3 domain in active Fyn. Another critical design consideration was to minimize any perturbation caused due to mCerulean insertion into Fyn. To this end, we selected a flexible poly-(glycine-serine) peptide linker (Trinh et al., 2004) to insert mCerulean within Fyn (also see Materials and method and Figure 3—figure supplement 1 for details on molecular engineering). We then examined the suitability of the designed biosensor constructs for cellular imaging. Immunoblotting analysis on cell lysates showed robust intracellular expression of full-length mCerulean-Fyn fusion proteins (hereby referred to as ‘mCer-Fyn’), confirming that the fusion proteins remain intact and resistant to proteolysis in cells (Figure 3—figure supplement 2A–I). Further, mCer-Fyn fusions expressed in cells showed expected fluorescence; with spectra from cells closely resembling unmodified mCerulean (Figure 3—figure supplement 2C). Additionally, the mVenus-F29 fusion biosensor protein also expressed well, was resistant to proteolysis (Figure 3—figure supplement 3A) and showed the expected fluorescent signatures resembling unmodified mVenus (Figure 3—figure supplement 3B). Importantly, we then tested if labeled Fyn kinase retains activity and is regulated appropriately. Src family kinases possess autocatalytic activity; a tyrosine residue (Y420 in h-Fyn) in the activation loop is phosphorylated when the protein gets activated (Cooper and MacAuley, 1988; Barker et al., 1995). Significantly, immunoblot analysis showed that mCer-Fyn retains autocatalytic activity similar to the wild-type unmodified protein in cells. Further quantification of Fyn Y-420 autophosphorylation levels, show no significant differences between mCer-Fyn and unlabeled wt-Fyn (Figure 3—figure supplement 2A,B, Figure 3—figure supplement 2—source data 1). Importantly, these data show that mCerulean insertion is minimally disruptive to kinase activity and that the engineered kinase appears to be regulated similar to unmodified Fyn.

To further test if our Fyn fusion (mCer-Fyn) is functional and can accurately reflect cellular Fyn dynamics, we examined if mCer-Fyn can modulate downstream signaling and functionally replace endogenous, wild-type Fyn. For this we carefully assayed Fyn effects on downstream signaling, as measured through extracellular signal-related kinase (ERK) phosphorylation and performed Fyn knockdown rescue in cells, comparing both unlabeled as well as Fyn-fusion (mCer-Fyn) constructs. We first established that RNA-i leads to a specific knockdown of cellular Fyn protein (Figure 3—figure supplement 4A) but not the closely related Src and Yes kinases in HEK293T cells (Figure 3—figure supplement 4B,C, Figure 3—figure supplement 4—source data 1). This Fyn knockdown significantly attenuates the levels of ERK phosphorylation in cells (Figure 3—figure supplement 5A,C, Figure 3—figure supplement 5—source data 1). This is consistent with earlier reports documenting Fyn’s role in modulating ERK activity and downstream signaling (Wary et al., 1998). Notably, expressing either mCer-Fyn or wt, unlabeled Fyn in ‘Fyn-knockdown’ cells significantly rescues this reduction in ERK phosphorylation. Further, the extent of increase in this ERK phosphorylation, mediated through ectopic Fyn expression in ‘Fyn-knockdown’ cells, was observed to be the same for wild-type, unlabeled Fyn as well as mCer-Fyn (Figure 3—figure supplement 5B–D, Figure 3—figure supplement 2—source data 1). These results unequivocally demonstrate that our labeled Fyn (mCer-Fyn) is regulated appropriately, ‘signaling competent’ and can functionally replace native, untagged Fyn; thereby making it suitable for faithfully reporting cellular Fyn dynamics.

Biosensor variants were then tested for efficacy in reporting intracellular Fyn activity. For this, we expressed biosensor constructs in HEK-293T cells and recorded fluorescence spectra from live cells (see Materials and methods). When cells co-expressing FRET donor (mCer-Fyn) and FRET acceptor (mVenus-F29) were excited at 435 nm (mCerulean/donor excitation), emission peaks were seen at ~475 nm (mCerulean) as well as at ~525 nm corresponding to the peak emission wavelength of the acceptor fluorophore (mVenus) (Figure 3D). Appearance of acceptor emission (mVenus,~525 nm) on donor excitation (435 nm) clearly shows that FRET occurs between mCer-Fyn and mVenus-F29. This confirms that mVenus-F29 can recognize intracellular Fyn in live cells and this recognition can be reported through a robust FRET signature. We then tested if this FRET signal is sensitive to the relative stoichiometry of the target (Fyn) and the binder F29 as would be expected with a binding-induced FRET. Indeed, increasing the amount of intracellular mVenus-F29 while maintaining constant expression of mCer-Fyn, led to a concomitant increase in FRET signal, followed by saturation of this FRET increase. Here, the saturation in FRET signal suggests that even increasing concentrations of F29 binder in cells does not cause an inadvertent or artefactual activation of Fyn kinase, especially since F29 binder expression in cells does not lead to any measurable changes in autocatalytic activity of Fyn (Y-420 autophosphorylation) (Figure 3—figure supplement 6, Figure 3—figure supplement 6—source data 1). Importantly, the FRET signal in cells is specific and dependent on mVenus-F29 binding to mCer-Fyn, since making a single point mutation in F29 abolishes the FRET response. If the non-binding variant of F29 (mVenus-F29-P41A) is co-expressed with mCer-Fyn in cells, little or no FRET is seen, even at higher concentrations of the non-binding control (Figure 3D left inset, Figure 3—source data 2). Further, the FRET signal was higher in cells co-expressing mVenus-F29 and constitutively active Fyn, relative to wild-type Fyn (Figure 3D right inset, Figure 3—source data 3). This shows that F29 binding and biosensor readout (FRET response) is indeed sensitive to the activation status of Fyn (Figure 3D).

Fyn kinase has two acylation marks (a myristoyl and a palmitoyl group) that make it preferentially localized to the cell membrane (van't Hof and Resh, 1999). Therefore, we reasoned that localization of F29 to the cell membrane would increase its proximity to membrane-bound Fyn, resulting in an increase in FRET signal and consequently greater dynamic range of the biosensor. To test this hypothesis, we added a myristoylation(‘myr’) sequence (MGSSKSKPKDPS) to the F29 binder (Victor and Cafiso, 1998). Indeed, addition of a myristoylation signal led to a substantial increase in FRET signal (Figure 3E, Figure 3—source data 4). To further test the fidelity of this enhanced FRET response observed with myristoylated-F29, we tested the non-binding variant (P41A) of the myr-mVenus-F29 binder. Notably, myristoylated non-binding mVenus-F29 P41A showed little or no FRET signal when co-expressed with mCer-Fyn (Figure 3E). Taken together, these results clearly show that the FRET signal observed with the F29-based Fyn biosensor is indeed due to specific binding of F29 to the active form of Fyn, and not just due to non-specific membrane localization and incidental proximity. The biosensor constructs –mCer-Fyn and myr-mVenus-F29 (referred to as binder) – are collectively referred to as FynSensor hereafter.

Src family kinases, including Fyn are known to strongly regulate cell adhesion as well as the cytoskeleton. Several SFK substrates and interacting partners are reported to be actin binding proteins, regulators of Rho family GTPases and other proteins that help remodel and regulate the actomyosin network (Ridley et al., 2003; Etienne-Manneville and Hall, 2001; Huveneers and Danen, 2009; Roca-Cusachs et al., 2012). We therefore reasoned that measuring the cell morphodynamics (Gulyani et al., 2011; Hodgson et al., 2016) would be a sensitive way to examine if moderate-to-low expression of labeled Fyn and the F29 binder significantly and measurably perturb the cell. First, the expression levels of FynSensor constructs labeled Fyn (mCer-Fyn) as well as mVenus-F29 were quantified in U2OS cells to ensure that the binder expression causes no artefactual activation of Fyn kinase (levels of pTyr416/total Fyn; Figure 3—figure supplement 6). We then quantitatively examined temporal changes in cell area and perimeter of cells expressing either FynSensor (mCer-Fyn:myr-mVenus-F29, DNA ratio 1:2, also see Figure 3D) or the binder alone or a control construct (myr-tagged-mVenus) in the adherent osteosarcoma U2OS cells at the expression levels specified. We find no significant change in cellular morphodynamics as a result of FynSensor expression the binder alone as compared to controls cells (Figure 3—figure supplement 7, Figure 3—figure supplement 7—source data 1). We further confirmed that the expression of Fyn-binder does not perturb the overall intracellular localization of labeled Fyn (Figure 3—figure supplements 8 and 9, Figure 3—figure supplement 9—source data 1). Immunofluorescence data also shows that labelled Fyn (mCer-Fyn) localization is similar to endogenous Fyn (Figure 3—figure supplement 9A).

While FynSensor and F29 expression had no effect on cell morphodynamics and Fyn localization, we also examined if FynSensor expression significantly perturbs downstream signaling. For this, we again used the extent of ERK phosphorylation as a sensitive readout of Fyn signaling. We find that neither expression of FynSensor nor binder alone in adherent osteosarcoma U2OS cells under the conditions used has any measurable effect on ERK phosphorylation levels (Figure 3—figure supplement 10, Figure 3—figure supplement 10—source data 1). Similarly, expression of FynSensor constructs (labeled Fyn + binder) in C2C12 mouse myoblasts (Figure 3—figure supplement 11, Figure 3—figure supplement 11—source data 1) or Fyn-knockdown HEK-293 cells did not significantly change levels of p-ERK (Figure 3—figure supplement 5). These data collectively show that FynSensor expression under these specified conditions does not significantly perturb downstream signaling. In light of these data showing little or no cellular perturbation, we have used the same FynSensor expression conditions for all further biosensor imaging studies.

FynSensor reports localized active Fyn in live cells

Our results show that FynSensor produces a FRET signal upon specific binding of F29 to Fyn-SH3, resulting in a sensitive readout of active intracellular Fyn. We further evaluated FynSensor for its ability to report spatial patterns of active Fyn in single, living cells. To investigate the spatial FRET response generated by FynSensor, we first employed the ‘fluorescence recovery after acceptor photobleaching (APB)’ method. APB analysis is a robust method of confirming and quantifying FRET in cells (Karpova and McNally, 2006), where recovery of donor fluorescence on APB is seen as a strong indicator of proximity-induced FRET. Energy transfer from the donor to the acceptor molecules results in quenching of donor fluorescence, and this quenching is relieved when proximal acceptor molecules undergo photobleaching. Indeed, when adherent osteosarcoma U2OS cells expressing FynSensor were illuminated with increasing doses of 514 nm laser light (wavelength corresponding to acceptor absorption; mVenus photobleaching), we observed a substantial, dose-dependent recovery of donor (mCerulean) fluorescence. This confirms FRET between mCer-Fyn and mVenus-F29. Significantly, APB-induced recovery of donor fluorescence was greater at cell edges (Figure 4A,B, Figure 4—source data 1), suggesting spatial variation in Fyn activity. Notably, in control cells expressing a mutant form of labeled-F29 (P41A), acceptor photobleaching showed no such dose-dependent recovery of donor fluorescence (Figure 4C, Figure 4—source data 2). Since the P41A mutant lacks the ability to bind Fyn-SH3, these data show that APB-induced recovery of donor fluorescence and the FRET response are due to F29 binding to target and not due to incidental proximity or optical artifacts. Finally, when cells were treated with a known pharmacological inhibitor of Src family kinase including Fyn, SU6656 (Blake et al., 2000), we observed a significant reduction in the amount of donor fluorescence being recovered (Figure 4—figure supplement 1, Figure 4—figure supplement 1—source data 1). This reduction in FRET (F29 binding to Fyn) on inhibitor treatment is interesting and points to allosteric changes in Fyn conformation on inhibitor binding. While ATP-competitive inhibitors like SU6656 directly bind the catalytic domain, they can also stabilize an inactive conformation of Src family kinases and modulate the accessibility of its key regulatory domains to ligands/binding proteins (Krishnamurty et al., 2013). Therefore, our data (reduced FynSensor FRET) suggests that indeed Fyn likely adopts a closed conformation in response to inhibitor treatment, thereby reducing SH3 domain accessibility/binding to F29. The sensitivity of FynSensor FRET to inhibitor binding in fact suggests that the labeled kinase shows allosteric conformational changes similar to those seen with untagged SFKs, further indicating that the biosensor faithfully reports Fyn conformational dynamics. Taken together, these results further confirm that the FynSensor readout is the result of specific binding of F29 binding to Fyn-SH3, and selectively reports the active form of Fyn (Figure 4B,C and Figure 4—figure supplement 1). The FynSensor readout also shows hitherto uncharacterized spatial patterns in intracellular active Fyn in live cells, with greater Fyn activity closer to the cell edge.

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*FynSensor* allows direct visualization of spatially localized active Fyn in cells.

Acceptor photo-bleaching (APB) experiment for visualizing active Fyn in U2OS cells co-expressing FRET donor (Fyn WT) and acceptor (myr-mVenus-F29) *FynSensor* constructs. (A) APB shows cellular FRET and localized Fyn activity. Representative figure shows donor and acceptor fluorescence images, prior to and after laser-based bleaching of acceptor with 514 nm laser pulse. Also shown is the corresponding ∆Donor image (see Materials and methods) (Scale bar = 10 μm). APB experiments reveal areas showing greater recovery of donor fluorescence after APB indicating spatially localized Fyn activity (B) APB-induced recovery of donor fluorescence is greater at the cell periphery and sensitive to bleaching light dosage. Percentage recovery of donor fluorescence in *FynSensor*-expressing cells, after acceptor photobleaching at different light dosage (using Equation 2, see Materials and methods), plotted as a function of distance from the cell edge. Extent of recovery also shown at different light dosage. Bleaching is carried out for 10 s using the 514 nm laser line at 2X, 5X and 10X doses (X = 0.069 mW). Lines represent mean of n = 14 cells for 10X, n = 9 cells for 5X, n = 7 cells for 2X, respectively. Values are mean ± s.e.m. (C) FRET signal measured through the APB method is specific (binding dependent) as well as sensitive to APB light doses. Cumulative APB-induced percentage recovery of donor fluorescence observed at the cell periphery in *FynSensor*-expressing cells (as in B), plotted as a function of light dosage (solid bars). Also, shown extent of recovery when donor-labeled Fyn is used with acceptor-labeled non-binding P41A mutant of F29 (*dashed bars*). Extent of recovery with non-binding control is minimal and significantly lower than observed for F29 *FynSensor.* Bars represent mean of n = 14 cells for 10X, n = 9 cells for 5X and n = 7 cells for 2X when binder/F29 is used and n = 14 cells for 10X, n = 10 cells for 5X and n = 8 cells for 2X when non-binder/F29P41A is used, respectively. Student’s unpaired one-tailed t-test was used to determine the p-value. Graph was made using GraphPad Prism. Values are mean ± s.e.m.

Figure 4—source data 1 Light dosage-dependence of donor fluorescence recovery in acceptor photo-bleaching experiment with F29 biosensor.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig4-data1-v1.xlsx
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Figure 4—source data 2 Light dosage-dependence of donor fluorescence recovery in acceptor photo-bleaching experiment with F29 biosensor and non-binding mutant.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig4-data2-v1.xlsx
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FynSensor reveals pulsatile Fyn activity and spatially compartmentalized signal integration

We used FynSensor to examine the spatial and temporal dynamics of Fyn activity, especially in light of the important role played by Fyn as a signaling node functioning downstream of distinct receptors classes. We reasoned that ability to specifically image the active conformation of a single SFK would provide new insight into how signal integration may occur. We first examined Fyn activation dynamics in serum-starved cells plated on fibronectin (FN) that show constitutive protrusion retraction cycles. For imaging activity dynamics, we used ratiometric sensitized emission (normalized acceptor emission due to donor excitation) to quantify FRET signals from the sensor (Gordon et al., 1998) (Materials and methods). Ratiometric sensitized emission measurements allow facile and rapid imaging of FRET responses in cells with both spatial and temporal resolution.

Sensitized emission (FRET) imaging with FynSensor in FN-plated serum-starved U2OS cells revealed spatially localized Fyn activity, with clear intracellular zones showing higher total FRET (FRET_T). Such intracellular zones showing enhanced FynSensor FRET_T were consistently observed in all cells in the data set (Figure 5A–I). Since all cells spontaneously appear to show regions of differential Fyn activity, we employed quantitative image analysis to probe these patterns of Fyn activity. An automated cell quadrant analysis of intracellular FRET_T levels provides clear evidence of differential and compartmentalized Fyn activity. Briefly, every cell imaged was divided into quadrants in an automated manner and FRET response for each quadrant was analyzed over time. These analyses show substantial and significant differences in Fyn activation levels (FynSensor FRET_T) between intracellular quadrants (designated as ‘low’ and ‘high’ FRET quadrants for consistency and comparison) (Figure 5A–II, Figure 5—source data 1). Importantly, the spatial patterns of Fyn activation revealed through FynSensor imaging are robust and are unlikely to be an artifact of imaging, since a single point mutation in the binder substantially abolishes the FRET response seen. When mCer-Fyn is co-expressed with the non-binding P41A mutant of F29, little or no FRET signal is seen in single cells, clearly showing that FynSensor FRET is dependent on F29 binding to active Fyn (Figure 5B, Figure 5—source data 2, Figure 5—figure supplement 1, Figure 5—figure supplement 1—source data 1, Figure 5—video 1). Overall, the spontaneous compartmentalizing of Fyn activity was observed in all cells and is a striking finding revealed through FynSensor imaging. (Figure 5A).

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*FynSensor* reveals integrin-dependent Fyn activity to be spatially compartmentalized.

(A) Spontaneous formation of zones enriched in active Fyn is seen in serum starved cells plated on fibronectin (FN). (I) Representative confocal fluorescence micrographs showing sensitized emission (total FRET: FRET_T) levels indicative of active Fyn in serum starved-U2OS cells expressing *FynSensor* (Scale bar = 10 μm). For quantitative image analysis of compartmentalized Fyn activity, we have divided the cell into quadrants labelled as Q1-Q4 as shown. (II) Bar graph comparing *FynSensor* FRET levels in distinct cellular zones. The mean of Max-FRET_T-HFQ (HFQ: high-FRET quadrant, Q4 in panel I) and Max-FRET_T-LFQ (LFQ: low-FRET quadrant, Q2 in I) values (see Materials and methods) are plotted. Values are mean ± s.e.m. Student’s paired one-tailed t-test has been used to determine the p-value (n = 37 cells). (B) *FynSensor* FRET readout is dependent on F29 binding Fyn, with the non-binding control mutant of F29 (P41A) showing no or significantly reduced FRET_T signal compared to *FynSensor*. (I) Confocal fluorescence micrographs showing sensitized emission (total FRET: FRET_T) levels for cells expressing mCer-Fyn and non-binding control binder mVenus-F29 P41A. (II) Comparison of FRET_T levels for *FynSensor* with non-binding control. The bar graph shows the average maximum FRET_T obtained (Max-FRET_T-HFQ) for non-binder (n = 5 cells) and *FynSensor* (n = 37 cells). Values are mean ± s.e.m. Student’s unpaired one tailed-t-test was used to determine the p-value. (C) Fyn activity levels are sensitive to inhibition of focal adhesion kinase (FAK), a known mediator of integrin signaling. (I) Confocal fluorescence micrographs showing *FynSensor* FRET_T levels before and after treatment with 10 μM of FAK inhibitor (PF 562271). Loss of basal Fyn activity is observed in cells when treated with inhibitor but not with DMSO (vehicle control). (II) Time course of Fyn activity in response to FAK inhibition. Average fluorescence intensity profile (FRET_T) of *FynSensor* cells (n = 5) over time, before and after addition of FAK inhibitor. (Scale bar = 10 μm). (III) Quantifying *FynSensor* FRET on FAK inhibition. Bar graph shows the average FRET_T intensity of cells before FAK inhibitor treatment (-FAKi), five mins after FAKi treatment and 15 mins after FAKi treatment (n = 5 cells). Values are mean ± s.e.m. Student’s paired one tailed-t-test was used to determine the p-value. (IV) FAK-inhibitor induced reduction in *FynSensor* FRET levels is NOT seen with vehicle control. Bar graph shows time-averaged, mean cell FRET_T intensity before and after treatment with the vehicle control (n = 8 cells). Values are mean ± s.e.m. Student’s paired one tailed-t-test was used to determine the p-value.

Figure 5—source data 1 Quantification of FynSensor FRET levels in low and high activity zones in U2OS cells.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig5-data1-v1.xlsx
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Figure 5—source data 2 Quantification of FynSensor non-binding control FRET levels in low and high activity zones in U2OS cells.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig5-data2-v1.xlsx
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Figure 5—source data 3 Quantification of FynSensor FRET levels in cells in treated with FAK inhibitor.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig5-data3-v1.xlsx
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We demonstrate that the spatially enhanced FRET_T patterns observed are unlikely to be an artifact of an unequal or spatially-constrained intracellular distribution of the F29 binder. Figure 5—figure supplement 2 (Figure 5—figure supplement 2—source data 1) shows that while the FynSensor FRET levels are non-uniform and spatio-temporally patterned, the fluorescently labeled F29 (myr-mVenus-F29) is homogeneously distributed in cells. While increased recruitment of Fyn may contribute to the establishment of regions of ‘high Fyn activity’ in serum-starved, FN-plated cells, our data analysis shows that differential Fyn activity observed cannot solely be ascribed to increased localization of Fyn. Figure 5—figure supplement 3 (Figure 5—figure supplement 3—source data 1) show that intracellular regions showing higher FRET_T also show significantly higher levels of donor-normalized FRET (donor-normalized FRET = FRET_T/mCer-Fyn fluorescence), which normalizes the overall activity for protein levels. This is also illustrated (Figure 5—figure supplement 4, Figure 5—figure supplement 4—source data 1) by examples of regions that have almost similar levels of the kinase (mCer-Fyn localization as visualized in the donor channel in regions marked 1 and 2) but show significantly different levels of kinase activity (FRET index image, regions 1 and 2), reaffirming the point that increases in FRET signals are just not due to increase in kinase localization.

We surmised that spatially enhanced Fyn activity seen in serum-starved cells on fibronectin may be caused by differential integrin signaling. To test if this Fyn activation in serum-starved cells, as visualized through FynSensor, is indeed sensitive to integrin signaling, we treated cells with an inhibitor of focal adhesion kinase (FAK), PF-562271 (Mills et al., 2015; Stokes et al., 2011) and examined FynSensor response (Figure 5C–I, Figure 5—video 2). Focal adhesion kinase (FAK) is a critical transducer of integrin signaling and is known to partner SFKs in mediating cellular responses (Renshaw et al., 1999; Cheng et al., 2014). Strikingly, when serum-starved, FN-plated U2OS cells were treated with PF-562271, there was an immediate and drastic reduction in the FynSensor FRET response (Figure 5C II-IV, Figure 5—source data 3). This reduction in activity is accompanied by a substantial dampening of the temporal changes seen in levels of active Fyn in cells (see below). These results clearly demonstrate that localized Fyn activity, reported through FynSensor, is sensitive to focal adhesion kinase activity; and supports the model that compartmentalized Fyn activity arises through differential integrin signaling in serum-starved adherent cells.

To confirm that these spatial patterns of Fyn activity are not just limited to any one specific cell-type, we performed FynSensor imaging in mouse C2C12 myoblasts apart from U2OS osteosarcoma cells. Notably, FynSensor again shows Fyn activity to be spatially patterned in the serum-starved C2C12 myoblast cells (Figure 6A). Quantitative quadrant analysis confirms differential Fyn activity (FRET_T) in distinct intracellular zones (Figure 6A–II, Figure 6—source data 1). To assess if cells with moderately overexpressed levels of Fyn kinase also show similar spatial-activation patterns, we performed imaging experiments in HEK-293T cells where the endogenous Fyn has been depleted through RNA-i and then complemented with FynSensor (Figure 3—figure supplement 9, Figure 6—figure supplement 1, Figure 6—figure supplement 1—source data 1). Biosensor imaging in these HEK cells indeed showed activation patterns very similar to those previously observed in U2OS and C2C12 cells (Figure 6B, Figure 6—source data 2). The observation of similar, spatially enhanced patterns of Fyn activity in these very distinct cell types shows that FynSensor responses are highly robust and is suggestive of conserved spatially modulated signaling mechanisms.

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Conserved spatial pattern of Fyn activity in distinct cell-types.

(A) *FynSensor* in mouse myoblast C2C12 cells reveals spontaneously compartmentalized Fyn activity, similar to that observed in U2OS cells. (I) Representative confocal fluorescence micrographs of serum-starved cells showing intracellular zones enriched with active Fyn when plated on FN (Scale bar = 10 μm). Quantitative image analysis, as previously described, yielded zones of high FRET_T and low FRET_T. (II) Bar graph comparing *FynSensor* FRET levels in distinct cellular zones. Graph compares the mean of Max-FRET_T-HFQ and Max-FRET_T-LFQ in serum-starved cells. Values are mean ± s.e.m. Student’s paired one-tailed t-test has been used to determine the p-value (n = 32 cells). (B) *FynSensor* imaging reveals spatially localized Fyn activity patterns when *FynSensor* is ectopically expressed in HEK293T Fyn-knockdown cells. (I) Confocal micrographs show zones of high and low FRET_T, showing spatially localized Fyn activity, similar to that seen with U2OS and C2C12 myoblasts. (II) Bar graphs comparing the mean of Max-FRET_T-HFQ and Max-FRET_T-LFQ in serum-starved cells confirm differential FRET levels across intracellular zones. Values are mean ± s.e.m. Student’s paired one-tailed t-test has been used to determine the p-value (n = 24 cells). (C) Spatial patterns of locally enhanced Fyn activity revealed by *FynSensor* imaging are unaffected by *FynSensor* expression levels. *FynSensor* imaging reveals locally enhanced, compartmentalized Fyn activity independent of sensor expression levels. To assess the effect of kinase expression on the FRET patterns observed in all the distinct cell types, we quantified the donor (m-Cerulean) fluorescence intensity for all the cells used in our study. Scatter-plot comparing donor fluorescence across different cell types, n = 32 cells for C2C12, 37 for U2OS and 24 for HEK-KD cells. Mean ± s.e.m is denoted. Student’s unpaired one-tailed t-test has been used to determine the p-value. Representative cells from the U2OS data set, having either relatively ‘high’ (above the mean value of 51.61 ± 1.805 fluorescence intensity units, denoted by *teal* box) or ‘low’ (below the mean value, *teal* box) *FynSensor* expression (donor/m-Cer fluorescence intensities), show very similar spatially compartmentalized Fyn activity pattern, suggesting that the activity patterns observed in different cells are not subject to expression artifacts.

Figure 6—source data 1 Quantification of FynSensor FRET levels in low- and high-activity zones in C2C12 cells.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig6-data1-v1.xlsx
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Figure 6—source data 2 Quantification of FynSensor FRET levels in low- and high-activity zones in Fyn-KD HEK293T cells.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig6-data2-v1.xlsx
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Figure 6—source data 3 Quantification of expression of FynSensor-Fyn (mCer-Fyn) in different cell-types showing similar activity patterns.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig6-data3-v1.xlsx
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To further confirm that the conserved patterns of Fyn activity are not impacted by changes in FynSensor expression levels, we systematically examined FynSensor FRET profiles as a function of expression. In all three cell types examined, the spatial patterns remain conserved independent of the precise levels of labeled Fyn (mCer-Fyn) as well as the binder (mVenus-F29) (Figure 6C, Figure 6—source data 3, Figure 6—figure supplement 2, Figure 6—figure supplement 2—source data 1 and Figure 6—figure supplement 2—source data 2).

Another consistent and striking feature revealed by FynSensor is the temporal bursts of Fyn activity. FRET_T levels show clear oscillations over time in serum-starved cells plated on FN and imaged (Figure 7A, Figure 7—source data 1, Figure 7—figure supplement 1, Figure 7—figure supplement 1—source data 1, Figure 7—video 1). Interestingly, these oscillations in Fyn activity revealed through FynSensor imaging are seen in multiple cell types and appears to be a conserved feature of Fyn activation dynamics. FRET_T profiles show temporal oscillations in C2C12 myoblast (Figure 7B, Figure 7—source data 2, Figure 7—figure supplement 2, Figure 7—figure supplement 2—source data 1), Fyn-KD HEK-293T (Figure 7C, Figure 7—source data 3, Figure 7—figure supplement 3, Figure 7—figure supplement 3—source data 1) as well as U2OS cells (Figure 7A) expressing FynSensor. We also tested if these Fyn activity pulses arise due to any sudden changes in kinase localization. Figure 5—figure supplement 3B–I clearly shows that along with the FynSensor FRET_T profiles, even the donor/mCerulean normalized FRET levels show pulsatile behavior. This shows that pulses observed are not due to fluctuations in Fyn concentration levels but arise due to direct and rapid regulation/modulation of Fyn activity (Figure 5—figure supplement 3B, Figure 5—figure supplement 3—source data 2). Since the pulsatile nature of Fyn activity was found to be conserved across cell types, we specifically examined the duration/frequency of these pulses in U2OS, C2C12 and HEK-Fyn KD cells. Figure 7D, Figure 7—source data 4 shows the mean of the dominant time-period of FynSensor FRET pulses in all three cell-types, determined through power-spectrum density (PSD) analysis performed on the FRET_T time-traces. The PSD analysis reveals the mean of dominant time-period to be ~3.5 min in three very distinct cell types, again highlighting a conserved feature of Fyn activity dynamics. There is increasing evidence to suggest that signaling modules, including growth factor responses in cells may show pulsatile behavior (Coster et al., 2017; Warmflash et al., 2012; Weber et al., 2010). Bursts of activity are expected to bear signatures of complex positive and negative (inhibitory) feedback loops that are integral part of growth factor and other signaling modules (Avraham and Yarden, 2011; Sparta et al., 2015; Albeck et al., 2013). Interestingly, temporal oscillations are highly pronounced in regions of spatially-enhanced Fyn activity as can be seen by plotting FynSensor FRET_T against time as well as distance from the celledge. The time-distance 3-D plot of Fyn activity shows bursts of activity in a tight intracellular zone proximal to the edge. (Figure 7E–I,II, Figure 7—source data 5).

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*FynSensor* imaging reveals spatially-compartmentalized temporal bursts of kinase activity across distinct cell types.

(A) A striking feature revealed by *FynSensor* is the pulsatile nature of Fyn activation. (I) Representative confocal micrographs of a U2OS cell at different time points after being plated on FN, as indicated, reveal significant oscillations in the FRET_T intensity (area marked by *red box*) (Scale bar = 10 μm). (II) Time-traces of the FRET_T signal showing temporal pulses of Fyn activity. (B) *FynSensor* imaging in C2C12 mouse myoblast cells, a cell-type distinct from the U2OS human osteosarcoma cells, also show an oscillating Fyn activity profile. (I) Confocal micrographs from a C2C12 cell transiently expressing *FynSensor*, at different time-points post FN plating, shows temporal bursts of kinase activity (area marked by *red box*). (II) Time-trace showing the pulsatile temporal pattern in FRET_T signal in the cell shown. (C) *FynSensor* imaging reveals pulsatile activity patterns in Fyn knockdown cells that have been ‘rescued’ with labeled Fyn (mCer-Fyn). Fyn activity pulses are similar to those observed in U2OS and C2C12, showing temporal patterns are robust and conserved. (I) Representative cell confocal micrograph of RNAi-Fyn knockdown HEK-293 (KD) cells expressing *FynSensor* show temporal oscillations in FRET_T levels (area marked by *red box*). (II) Time-trace of FRET_T signal from individual cell revealing pulsatile Fyn signal. (D) Oscillatory/pulsatile behavior in levels of active Fyn in the different cell-types studied reveal a consistent major time period. Plotted here is the mean of the dominant time-period of *FynSensor* FRET pulses in U2OS C2C12 and HEK-KD cells expressing the sensor, determined through power spectrum density (PSD) analysis on the time-traces of single-cell quadrant averaged total FRET. The PSD analysis reveals the mean of dominant time-period to be ~3.5 min in distinct cell types. N = 37 for U2OS, 32 for C2C12 and 16 for HEK-KD cells. Values are mean ± s.e.m. Student’s two-tailed unpaired t-test has been used to determine the p-value. See Materials and methods for details of the analysis. (E) Pulsatile Fyn activity is spatially constrained. Intracellular zones of elevated Fyn activity also show clear oscillations over time. (I) Confocal fluorescence micrograph of a C2C12 cell shows a zone of high FRET signal (indicated by *yellow* box). For plotting temporal oscillations over distance from edge, an arc (of arbitrary length units) was suspended on the membrane encompassing this zone. A smaller sector was then created on the cell (indicated by *green* lines). (II) Plot shows a heat map of FRET_T intensity values in this sector, over time, as a function of distance from the cell- membrane. Data reveals that active kinase to be spatially constrained and oscillating over time. See Materials and methods for details of the analysis.

Figure 7—source data 1 Analysis of temporal patterns of Fyn activity in U2OS cell.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig7-data1-v1.xlsx
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Figure 7—source data 2 Analysis of temporal patterns of Fyn activity in C2C12 cell.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig7-data2-v1.xlsx
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Figure 7—source data 3 Analysis of temporal patterns of Fyn activity in Fyn-KD HEK 293 T cell.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig7-data3-v1.xlsx
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Figure 7—source data 4 Quantification of Fyn activity pulse time-period across different cell-types.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig7-data4-v1.xlsx
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Figure 7—source data 5 Quantification of Fyn activity relative to distance from cell membrane in C2C12 cell.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig7-data5-v1.xlsx
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Fyn kinase signals downstream of multiple receptor classes, including integrin and receptor tyrosine kinases (RTKs/Growth factor receptors). Based on prior evidence (DeRita et al., 2017; Edick et al., 2007; Samayawardhena et al., 2007; Parsons and Parsons, 1997), we hypothesized that Fyn, functioning as a signaling node may be involved in dynamically integrating signals downstream of integrins and RTKs (Lehembre et al., 2008; Kinnunen et al., 1998; Arias-Salgado et al., 2005). We used the FynSensor to investigate this putative signal integration. For this, we examined the effect of platelet-derived growth factor (PDGF) on FN-plated, serum-starved U2OS cells and visualized active Fyn using FynSensor FRET. As discussed earlier, serum-starved FN-plated cells already showed spatially enhanced and temporally regulated Fyn activity, consistent with constitutive cell polarization and Fyn activation downstream of spatially regulated integrin signaling (Figure 5A). However, after stimulation with PDGF, FynSensor cells (but not the P41A non-binding control Figure 8—video 1) showed a significant increase in Fyn activity as indicated by enhanced FRET signal (Figure 8A). Strikingly, despite a global PDGF stimulation, the increase in Fyn activity observed was spatially localized. It appeared that on stimulation, the FRET_T signal, preferentially increased in regions that already showed higher FRET in un-stimulated cells (Figure 8A; Figure 8—video 1, resulting in highly compartmentalized Fyn activity patterns.

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Integrin-dependent, spatially compartmentalized, pulsatile Fyn activity can be modulated by growth factor suggesting signaling crosstalk.

(A) Spontaneously formed zones of active Fyn in serum-starved cells plated on FN can be further modulated by Platelet-derived growth factor (PDGF). Representative confocal fluorescence micrographs showing sensitized emission (FRET_T) levels indicative of active Fyn in serum-starved (I) and PDGF-stimulated U2OS cells (II) (Scale bar = 10 μm). (B) Quadrant analysis of cells reveal that on a global PDGF stimulation of serum-starved cells, the increase in Fyn activity observed was spatially localized (I) Bar graph comparing the mean of Max-FRET_T-HFQ before and after addition of PDGF (10 ng/ml) indicates the FRET_T intensity increase in the high FRET zone on addition of PDGF. Values are mean ± s.e.m. Student’s paired one-tailed t-test has been used to determine the p-value (n = 18 cells). (II) PDGF-induced enhancement in Fyn activity is greater in pre-activated intracellular zones. Bar graph show the difference in the mean intensities of the maximum FRET_T (Max-FRET_T) obtained before and after PDGF addition for both ‘high FRET’ as well as ‘low FRET’ quadrants/zones across single cells. Quadrants were designated as ‘HFQ’ or ‘LFQ’ in FN-plated cells, prior to PDGF stimulation (n = 18 cells). Values are mean ± s.e.m. Student’s paired one-tailed t-test has been used to determine the p-value. (C) Focal adhesion kinase (FAK) activity not only regulates Fyn activity but is required for the modulation/enhancement of Fyn activity by PDGF. (I) Confocal fluorescence micrographs showing sensitized emission (FRET_T) levels in untreated/serum starved cells, after treatment with FAK inhibitor (FAKi, PF-562271 and then followed by further treatment with PDGF. Scale bar = 10 µm. FAK inhibition attenuates FRET levels (active Fyn levels) in FN-plated serum-starved cells AND abolishes the enhancement and modulation of Fyn activity by PDGF. FAKi treated cells fail to respond to PDGF as measured through *FynSensor* FRET levels. (II) The mean FRET_T intensity profile for the high FRET_T quadrant in 5 cells plotted over time. Also, marked in color are experimental condition or the treatments; serum starved cells (yellow, t = 0–10 mins), + FAKi (cyan, t = 10–20 mins) AND +FAKi+PDGF (lavender, t = 20–30 mins). Values are mean ± s.e.m. (III) Whole cell FRET_T data from all the cells (n = 5) for untreated cells, +FAKi, and +FAKi+PDGF. The mean intensity value of untreated cells has been used to normalize the data. Student’s two-tailed paired t-test has been used to determine the p-value. Values are mean ± s.e.m. Data show no significant enhancement is seen in Fyn activity by PDGF stimulation, if cells are pre-treated with FAKi.

Figure 8—source data 1 Quantification of PDGF-induced enhancement in Fyn activity across different cellular zones.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig8-data1-v1.xlsx
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Figure 8—source data 2 Quantification of FRET_T levels in untreated cells, followed by FAK inhibitor treatment and further treatment with PDGF.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig8-data2-v1.xlsx
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To quantitatively examine these activity patterns and signal modulation, we again employed automated image analysis. An automated intracellular quadrant analysis confirms differential FynSensor FRET levels across distinct intracellular zones, post-PDGF stimulation (Figure 8B–I,II, Figure 8—source data 1). These data demonstrate that cells remain ‘polarized’ with respect to Fyn activity despite a global PDGF stimulation. Notably, quadrant image analysis also confirmed a preferential PDGF-induced enhancement of Fyn activity in pre-activated intracellular zones. Figure 8B–II shows that PDGF-induced enhancement in FynSensor FRET is greater in intracellular quadrants already showing higher FRET_T pre-stimulation. Overall, this is a powerful demonstration of the ability of FynSensor to reveal hitherto unexplored aspects of Fyn signaling dynamics.

FynSensor shows Fyn activity to be compartmentalized even in serum-starved cells. This is likely due to differential integrin signaling, since Fyn activity is significantly attenuated upon FAK inhibition. This spatially constrained Fyn activity could be further increased through growth factor stimulation, with greater enhancement seen in pre-activated areas despite a global stimulation. These results provide a direct illustration of dynamic and intracellular localized signaling crosstalk between integrins and growth factor receptors; visualized through the activation (conformational change) of a non-receptor tyrosine kinase that get activated by each of these receptor classes. While results so far imply that differential integrin signaling appears to spatially restrict the effect of growth factor, we asked if appropriate integrin signaling is required for growth factor mediated modulation of Fyn activity. For this, we inhibited focal adhesion kinase activity and specifically tested if growth factor is still able to modulate Fyn activity levels. Interestingly, when FynSensor U2OS expressing cells are treated with PF-562271 (FAK inhibitor), not only are the FynSensor FRET levels drastically reduced, the cells become insensitive to growth factor stimulation as measured through Fyn activity. PDGF-stimulation of inhibitor treated cells show little or no change in FynSensor FRET levels (Figure 8C, Figure 8—source data 2, Figure 8—figure supplement 1, Figure 8—figure supplement 1—source data 1, Figure 8—video 2). These data indeed confirm a robust, spatio-temporally modulated and functional crosstalk between integrin and growth factor signaling.

FynSensor reveals striking spatio-temporal patterns of Fyn activity. Since Fyn is a key regulator of cell physiology, including cytoskeleton remodeling and adhesion dynamics, we also asked how Fyn activity correlates with membrane motility. For this we acquired images at faster acquisition speeds (~35 s/frame) and performed quantitative image analysis using the Fiji plugin ADAPT (Barry et al., 2015) and measured both membrane motility and FRET_T along the periphery of the cell, over the timecourse of the experiment (Figure 9A I-II, Figure 9—source data 1). Intriguingly, a plot of membrane motility versus FRET_T levels shows that the two parameters tend to be inversely correlated (Figure 9B). This is an interesting trend suggesting that intracellular regions showing higher Fyn activity is likely to show reduced overall membrane motility changes. This link between Fyn activity and dampened cell membrane oscillations points to a ‘poised’ cell membrane when/where Fyn is active. Such a notion is consistent with Fyn’s role in regulating cell-matrix adhesions and may need to be probed further (Figure 9C). It is remarkable that visualizing the conformational dynamics of a membrane-bound, non-receptor kinase reveals pulsatile patterns, which can be modulated through integrin and growth-factor signaling.

Figure 9

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Fyn activity couples to cellular morphodynamics.

(A) Fyn activity correlates with reduced amplitudes of membrane protrusion-retraction as revealed by simultaneous measurement of membrane motility and *FynSensor* FRET_T. Quantitative image analysis was performed using the ADAPT plugin [J Cell Biol (2015) 209 (1): 163–180]. (I) Representative 3D plots correlating Fyn activity (left) and membrane motility (right) for a single U2OS cell. Area within the dotted box marks region of high FRET_T and low membrane motility. (II) A representative confocal micrograph (FRET_T image) of the same U2OS cell showing Fyn activity to be spatially constrained (*Inset*: FRET channel image for clearly depicting the whole cell). (B) Graph shows the net membrane motility changes occurring over time, plotted against FRET_T intensity for U2OS cells (n = 15 cells). FRET_T levels are inversely correlated to membrane motility (Plot fitted using the equation f = y0+a*x+b*x^2+c*x^3; R (2) value = 0.8509). Values are mean ± s.e.m. (C) An illustration of Fyn activity being negatively correlated to membrane dynamics wherein, cellular zones showing higher Fyn activity (high FRET_T) show reduced overall changes in membrane motility (lesser constitutive protrusion/retraction cycles).

Figure 9—source data 1 Quantification of correlation between membrane motility and Fyn activity across multiple U2OS cells.: https://cdn.elifesciences.org/articles/50571/elife-50571-fig9-data1-v1.xlsx
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Discussion

Direct visualization of protein activity is essential in order to gain a quantitative understanding of dynamic signaling networks that govern cell behavior. Despite the critical roles played by Src family kinases (SFKs) in regulating physiology (Chetty et al., 2015; Gujral et al., 2014; Kim et al., 2009; Lewis-Tuffin et al., 2015; Nygaard et al., 2014; Saad, 2009; Timpson et al., 2001; Zhang et al., 2013; Zhang et al., 2014), specific tools/sensors to image activity of individual kinases in live cells and tissues are not available. This is particularly important as individual SFKs can perform overlapping but specific, even seemingly opposing roles, to control cellular output (Zhang et al., 2014; Kuo et al., 2005; Lowe et al., 1993; Lowell and Soriano, 1996; Marchetti et al., 1998; Molina et al., 1992; Palacios-Moreno et al., 2015). Therefore, understanding the specific activity patterns of individual kinases assumes considerable importance. Our work clearly addresses this issue and establishes a platform for developing new biosensors for visualizing activation of individual Src kinases. Using combinatorial library screening and protein engineering, we develop a biosensor for the critical SFK, Fyn. This specific biosensor reports the activation dynamics of Fyn in live cells, with no interference from other kinases. Fyn kinase is a major regulator of multiple cellular processes and has emerged as a key player in various disease pathologies (Nygaard et al., 2014; Bhaskar et al., 2005; Schenone et al., 2011; Chin et al., 2005), but this is the first direct visualization of Fyn activity in cells. Interestingly, currently there are no tools for imaging active Fyn, with even specific antibodies not being available.

Our approach is extremely general and can be used for building sensors for other Src family kinases that are critical players in homeostasis and diseases. In this light, our work on addressing the specificity of biosensor binding using NMR structural analysis is likely to be significant. While our targeted screening and binding analysis yields a new binder (F29) that is highly specific for Fyn (Figure 1), the NMR structure of F29 bound to Fyn SH3 provides fascinating new insights into the molecular basis of specificity (Figure 2, Figure 2—figure supplement 1). We identify key interactions that mediate binding and further use this insight to switch specificity of binding (Figure 2). For instance, our Fyn-specific binder F29 does not or weakly bind Src-SH3 (Figure 1C,E). However, based on predictions from structural analyses, making just a single residue change in Src SH3 allows it to now bind F29 better (Figure 2—figure supplement 3C). These findings offer the possibility of generating specific binders targeting various Src family kinases, leading to new biosensors and tools. More broadly, we showcase how targeted screening of combinatorial protein libraries can be deployed to generate genetically encoded biosensors for visualizing active conformations of signaling proteins. This is a generally applicable strategy for biosensor development. With this method, it is possible to develop sensors even for ‘difficult’ targets, where structural information is limited and conventional tools like antibodies do not exist.

We demonstrate that FynSensor is able to report on the activation status of Fyn in cells. Newly engineered binder F29 binds the active form of Fyn and this binding leads to a FRET response that can be measured in live cells (Figures 3–9). Our sensor design and extensive control experiments ensures high sensitivity while minimizing perturbation. We have shown that the fluorescently labeled Fyn retains kinase activity, is fully functional and behaves like the unmodified kinase in terms of its regulation, localization and ability to modulate downstream signaling (Figure 3—figure supplements 2 and 5–11). Importantly, the fluorescently labeled Fyn is also able to complement the endogenous Fyn in RNAi-knockdown rescue experiments. For instance, in knockdown-rescue experiments, labeled Fyn is able to rescue downstream ERK phosphorylation similar to untagged Fyn, showing that our tagging of Fyn does not perturb its regulation and signaling function. We also demonstrate that under our imaging conditions (low-to-moderate expression of FynSensor) in two very distinct cell types, biosensor does not perturb either downstream signaling or cellular morphodynamics, reiterating the efficacy of our design (Figure 3—figure supplements 6–11). When the non-binding control version of the F29 binder (F29P41A) is used in cells, the FRET response is abolished, showing FRET to be dependent on F29 recognizing activated Fyn (Figure 5B, Figure 5—figure supplement 1). Further, an inhibitor of kinase activity also significantly attenuates FRET response, showing the biosensor readout to reflect kinase activity (Figure 5C). Overall, we show that the FynSensor response is specific and reflects cellular activity of Fyn.

A significant advantage of FynSensor is that it is highly specific and directly reports on the active conformation of Fyn with high spatial and temporal precision, unlike previously reported kinase sensors. This direct visualization of the active form of a single kinase, within a critically important yet complex family of kinases, has led to new insights into signaling dynamics and regulation. FynSensor imaging reveals Fyn activity to be localized and temporally modulated (Figures 4, 5, 6, 7 and 8), with greater activity closer to the cell edge. Further, as serum-starved, fibronectin (FN)-plated cells undergo constitutive cycles of protrusion and retraction, we observe intracellular zones of high Fyn activity and even spontaneous cell polarization measured through levels of active Fyn. Critically, we show that this compartmentalized Fyn activity is dependent on integrin signaling. Inhibition of focal adhesion kinase, a key mediator of integrin signaling, abolishes or greatly attenuates Fyn activity. When FN-plated cells with spatially localized Fyn activity are treated with platelet-derived growth factor (PDGF), we observe a greater increase in Fyn activity in and around the zones that were already pre-activated. This is indeed remarkable. Despite global PDGF treatment, its effect is highly localized and is dependent on intracellular zones that are established through integrin signaling. Our results suggest a model wherein localized/differential integrin activation in cells undergoing constitutive protrusion-retraction cycles not only helps establish signaling zones/compartments with higher Fyn activity, but also sensitizes these zones to be more responsive to growth factor signaling (Figure 10). This could be through either preferential localization or pre-sensitization of PDGF-receptors and/or associated signaling components (Figure 10). Thus, compartmentalized integrin signaling is not only maintained but also ‘functionally enhanced’ through growth factor stimulation. We further show that integrin signaling, not only spatially constraints Fyn activity and subsequent growth factor response, but that appropriate integrin signaling is required for growth factor modulation of Fyn activity. Inhibiting focal adhesion kinase activity not only dramatically attenuates Fyn activity in FN-plated, serum-starved cells, it also renders Fyn insensitive to any further stimulation through PDGF (Figure 8C).

Figure 10

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Model representing how Fyn activity helps mediate spatio-temporally localized signaling crosstalk.

(A) Src kinases have been long known to associate with the integrin receptor and growth factor receptor (RTK) signaling, thereby placing these kinases at nodal spots for signal integration. We provide the first visual proof for Fyn's involvement in integrating signals from these two receptor classes and see that the cell is highly polarized with respect to Fyn kinase activity. *FynSensor* shows that integrin receptor engagement results in formation of ‘zones’ of active Fyn in the cell, causing the cell to become polarized with respect to the activity of the kinase, even in serum-starved, FN-plated cells. PDGF stimulation further increases Fyn activity preferentially in these pre-sensitized regions only, thereby strengthening the spatially localized response. (B) Spatio-temporally localized Fyn activity; specifically, set-up through signal integration offers a quantitative picture of how Fyn activity may help regulate cell fate in certain situations. Fyn is known to be crucial for context-dependent cell migration and cell state switching in normal as well malignant cells, therefore, a spatio-temporally polarized response may be important for setting up cell output, especially for processes like that of directional cell migration and adhesion dependent cell switching. Pulsatile Fyn activity further suggests pre-organized signaling clusters with negative and positive feedback loops sensitive to both integrin and growth factor signaling.

Our results offer a striking demonstration of spatially localized crosstalk between integrin and growth factor signaling and shows that integrin can localize and regulate the effect of even globally applied growth factors in activating downstream signaling (Figure 8). This illustrates how by visualizing the dynamic activation of a key signaling node (Fyn), it is possible to directly visualize signaling crosstalk between receptors in cells. Prior work has specifically implicated Fyn for its ability to integrate information emanating from different cell-surface receptors (Colognato et al., 2002; Colognato et al., 2004). For instance, Fyn is plays a crucial role in cell survival and myelination of neurons by oligodendrocytes, specifically by integrating integrin signaling with growth factor signaling. Oligodendrocyte precursors are able to survive and subsequently wrap neurons with a myelin sheath, even in low growth factor conditions through the additive effects of integrin and RTK signaling (Laursen et al., 2009; Laursen et al., 2011; Laursen and Ffrench-Constant, 2007; Sperber and McMorris, 2001; Schäfer et al., 2016). This signal integration is mediated through Fyn activity (Laursen et al., 2009; Schäfer et al., 2016). It is striking and highly significant that our biosensor reveals evidence of such signal integration, in single cell imaging experiments in-vitro. This shows that compartmentalized signaling crosstalk between different receptor classes may be a key feature of signaling systems (Figure 10).

Biosensor imaging also shows Fyn activity signals to be pulsatile. Recent work has shown that oscillatory signaling patterns are key hallmarks of several signaling modules, including regulation by transcription factors (Imayoshi et al., 2013; Isomura and Kageyama, 2014) as well as MAP kinases (Weber et al., 2010; Sparta et al., 2015; Albeck et al., 2013; Antoine-Bertrand et al., 2016; Shankaran et al., 2009). Generally, pulses arise due to the presence of negative feedback loops, which may also be accompanied by additional modulators. Interestingly, there is some evidence that switching temporal activity patterns can alter cellular fate; for instance, pulsatile ERK activity leads to one output while persistent activity leads to another. We observe robust and spontaneous activity pulses of Fyn in very distinct cell types even in serum-starved cells plated on FN (Figure 7), and these pulses are further modulated by growth factor stimulation. This behavior strongly suggests the presence of rapidly activated, negative feedback loops that directly modulate Fyn activity.

Fyn activity pulses may be functionally significant in light of its demonstrated role in regulating cell migration, adhesion and actomyosin remodeling. In migration, there is an intrinsic periodicity in cycles of actin polymerization/remodelling/depolymerisation, periods of membrane remodelling as well adhesion assembly and disassembly. A protein that regulates several of these processes is conceptually much more likely to have pulses of activity versus sustained activity. In this light, it is remarkable that we are able to pick up ‘activity pulses’ at specific locations in the cell. This would need to be explored further in the context of directed migration and control of cell proliferation. Further, its indeed striking that membrane motility is reduced in the regions/zones where Fyn is more active (Figure 9). Spatially-localized pulsatile activation may likely cause transient but limited stabilization of signaling clusters which would be difficult to achieve with sustained activation. These observations taken together suggest that Fyn activity may be correlated with a ‘poised’ membrane state and formation and stabilization of specific integrin-FAK dependent signaling clusters, which can be further tuned by growth factor signaling. Such specific control of adhesion and actomyosin dynamics may be important in Fyn’s ability to mediate directed cell migration, growth control and tissue invasion. Context dependent and spatio-temporally regulated Fyn activity may provide a new perspective on understanding Fyn’s roles in regulating diverse and sometimes opposing cellular functions. Fyn activity profiles also provide a direct readout of highly localized signaling crosstalk between distinct receptor classes. Overall, this work sets the stage for detailed investigations of such functional signaling crosstalk in numerous cellular contexts and offers a new paradigm for direct visualization of signaling dynamics mediated by Fyn and other Src kinases.

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High-throughput screening yields a new protein binder for generating a specific biosensor for active Fyn.

Figure 1—source data 1

Structure and binding interface of Fyn-SH3:F29 complex by NMR.

Developing a FRET biosensor for sensing active Fyn in cells.

Figure 3—source data 1

Figure 3—source data 2

Figure 3—source data 3

Figure 3—source data 4

FynSensor allows direct visualization of spatially localized active Fyn in cells.

Figure 4—source data 1

Figure 4—source data 2

FynSensor reveals integrin-dependent Fyn activity to be spatially compartmentalized.

Figure 5—source data 1

Figure 5—source data 2

Figure 5—source data 3

Conserved spatial pattern of Fyn activity in distinct cell-types.

Figure 6—source data 1

Figure 6—source data 2

Figure 6—source data 3

FynSensor imaging reveals spatially-compartmentalized temporal bursts of kinase activity across distinct cell types.

Figure 7—source data 1

Figure 7—source data 2

Figure 7—source data 3

Figure 7—source data 4

Figure 7—source data 5

Integrin-dependent, spatially compartmentalized, pulsatile Fyn activity can be modulated by growth factor suggesting signaling crosstalk.

Figure 8—source data 1

Figure 8—source data 2

Fyn activity couples to cellular morphodynamics.

Figure 9—source data 1

Model representing how Fyn activity helps mediate spatio-temporally localized signaling crosstalk.

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Ananya Mukherjee

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Randhir Singh

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Sreeram Udayan

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Sayan Biswas

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Pothula Purushotham Reddy

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Saumya Manmadhan

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Geen George

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Shilpa Kumar

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Ranabir Das

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Balaji M Rao

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Akash Gulyani

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