Yuhao Ji,Taniya Chakraborty,Seraphine V. Wegner
Self-Regulated and Bidirectional Communication in Synthetic Cell Communities
synthetic cells,adaptive,bidirectional communication,chemiluminescence,self-regulation
Cell-to-cell communication is not limited to a sender releasing a signaling molecule and a receiver perceiving it but is often self-regulated and bidirectional. Yet, in communities of synthetic cells, such features that render communication efficient and adaptive are missing. Here, we report the design and implementation of adaptive two-way signaling with lipid-vesicle-based synthetic cells. The first layer of self-regulation derives from coupling the temporal dynamics of the signal, H2O2, production in the sender to adhesions between sender and receiver cells. This way the receiver stays within the signaling range for the duration sender produces the signal and detaches once the signal fades. Specifically, H2O2 acts as both a forward signal and a regulator of the adhesions by activating photoswitchable proteins at the surface for the duration of the chemiluminescence. The second layer of self-regulation arises when the adhesions render the receiver permeable and trigger the release of a backward signal, resulting in bidirectional exchange. These design rules provide a concept for engineering multicellular systems with adaptive communication.
Self-Regulated and Bidirectional Communication in Synthetic Cell Communities

Cell-to-cell communication is not limited to a sender releasing a signaling molecule and a receiver perceiving it but is often self-regulated and bidirectional. Yet, in communities of synthetic cells, such features that render communication efficient and adaptive are missing. Here, we report the design and implementation of adaptive two-way signaling with lipid-vesicle-based synthetic cells. The first layer of self-regulation derives from coupling the temporal dynamics of the signal, H2O2, production in the sender to adhesions between sender and receiver cells. This way the receiver stays within the signaling range for the duration sender produces the signal and detaches once the signal fades. Specifically, H2O2 acts as both a forward signal and a regulator of the adhesions by activating photoswitchable proteins at the surface for the duration of the chemiluminescence. The second layer of self-regulation arises when the adhesions render the receiver permeable and trigger the release of a backward signal, resulting in bidirectional exchange. These design rules provide a concept for engineering multicellular systems with adaptive communication.

Cell-to-cell communication through soluble chemical signals determines how cells function in a community. The exchange between sender and receiver cells makes it possible to coordinate the behavior of individuals so that they function as one multicellular entity. This includes, but is not limited to, different bacteria-forming biofilms, interactions of microorganisms with their hosts, and cellular communication in multicellular organisms. Analogously, synthetic cells, assembled from molecular building blocks, can work together by implementing the basic principles that underlie intercellular communication. Through the exchange of chemically encoded messages, synthetic cells in communities can influence each other, split cascade reactions between different members, develop prey–predator relationships, perform decentralized computing, and influence living cells. Cellular communication is not limited to a sender cell secreting a chemical signaling molecule and a receiver cell responding to it but is often more sophisticated and dynamic. In many cases, the communication is bidirectional and regulated in space and time, a prime example of it being the adaptive immune systems that require the transient and dynamic interaction of different types of immune cells. Coupling the temporal dynamics of signal generation with the spatial proximity of the sender and receiver renders the communication adaptive and increases the efficacy of signal delivery because it increases the probability of a receiver being within the signaling range at the right time. For this reason, sender cells can actively recruit receivers when they start producing signaling molecules through chemotactic gradients or direct adhesions. In return, receiver cells can signal their presence and separate from the sender cells once the signal weakens. In communities of synthetic cells, the efficiency of communication and its modulation are of major importance considering the limited resources for producing the signal. In addition, there are only a few examples of bidirectional communication in synthetic cells. One way to increase efficacy is to convert and amplify the signal through enzymatic cascade reactions and allosteric regulators. A second possibility is to bring sender and receiver into spatial proximity and geometries, either with the help of external compartments, microfluidics, ultrasound, or optical tweezers or through external modulation with stimuli such as temperature, metabolic activity, or light. Finally, local release of signaling molecules and at adhesion sites is another way of controlling efficacy. Yet, none of these examples matches the self-regulated and bidirectional communication that living cell communities are built on. Here, we report adaptive two-way communication between two populations of giant unilamellar vesicle (GUV)-based synthetic cells. In particular, the temporal production of the signaling molecule, H2O2, in the sender GUV is coupled to the spatial proximity of the receiver GUV (Figure 1, Steps 1 and 2). To achieve this, we demonstrate that chemiluminescence activates light-responsive proteins, which act as adhesion molecules between the vesicles. When the sender and receiver GUVs come into contact, the receiver GUV becomes more permeable, leading to the release of a backward signal, Ca2+, from the receiver to the sender (Figure 1, Step 3). The vicinity of the sender and receiver is self-regulated through the production of H2O2, and the two synthetic cells separate once the sender no longer produces it (Figure 1, Step 4). This study therefore shows how coupling the production of the signaling molecule, chemiluminescence activated adhesions, and membrane permeability results in self-regulated and bidirectional cell-to-cell communication.

As a first step, we sought a way to connect the production of an intracellular signaling molecule to extracellular adhesions between sender and receiver GUVs, thereby temporally regulating their spatial proximity (Figure 2a). We chose as the signaling molecule hydrogen peroxide (H2O2) and as the GUV-GUV adhesion module the proteins iLID and Nano, which bind to each other under blue light and dissociate from each other in the dark. We previously demonstrated that mixtures of GUVs functionalized with either iLID or Nano at their outer surface adhered to each other under blue light and separated when the illumination was stopped. To connect the production of the H2O2 signal with the photoactivation of the iLID–Nano interaction, we envisioned using chemiluminescence rather than external blue light illumination.

To establish the photoactivation with chemiluminescence, we used the horseradish peroxidase (HRP)-catalyzed chemiluminescence reaction between luminol and H2O2. The resultant light has an emission maximum of around 450 nm, which spectrally overlaps with the absorbance peak of iLID of around 430–480 nm (Figure 2b). The number of photons detected from the chemiluminescence reaction (200 μM luminol, 500 μM H2O2, 2 μg/mL HRP) decreased significantly in the presence of iLID (20 μM) (Figure 2c), in particular in the part of the spectrum where iLID absorbs (Figure S1). The persistent low photon output over time in the presence of iLID was attributed to the fast reversion kinetics of iLID (dark reversion rate 3.5 × 10–2 s–1, t1/2 = 20 s) and the plateau after ca. 8 min to a photostationary state. Altogether, these results showed that iLID is able to absorb photons produced in the chemiluminescence reaction. Direct evidence for the photoactivation of iLID through chemiluminescence and its subsequent binding to Nano was provided by a competitive fluorescence polarization assay (Figure S2). In this assay, the SsrA peptide labeled with a TAMRA fluorophore (SsrA-TAMRA) binds to Nano, resulting in an increase in fluorescence polarization. Upon photoactivation, iLID binds to Nano and displaces SsrA-TAMRA from the complex, which is observed as a decrease in fluorescence polarization. When this assay was performed in the presence of the chemiluminescence reaction, the initially high fluorescence polarization decreased over the course of 20 min as a result of binding of iLID to Nano in the photoactivated state (Figure 2d). This value remained constant for the next 10 min, but gradually returned to the initial value as the photon output of the chemiluminescence reaction decreased and iLID reverted to the dark state (Figure 2c). Control experiments lacking one of the essential reagents of the chemiluminescence reaction (luminol, H2O2, or HRP) did not result in changes of fluorescence polarization. It was not possible to monitor the photoactivation of iLID directly through the decrease in the peak around 450–480 nm because the reagents of the chemiluminescence reaction absorbed in the same range. Overall, these findings showed that it is possible to photoactivate iLID by using chemiluminescence and that reaction kinetics of the chemiluminescence reaction correlate with the binding between iLID and Nano. As the next step, we localized the chemiluminescence reaction inside the sender GUV in order to trigger adhesions between iLID-decorated sender GUVs and Nano-decorated receiver GUVs (Figure 2a). To achieve this, we encapsulated the catalyst, HRP (2 μg/mL), inside GUVs (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) with 10 mol % 1-palmitoyl-2-oleoyl-snglycero-3-phospho-(1′-rac-glycerol) (POPG)). Upon external addition of the membrane-permeable substrates luminol (200 μM) and H2O2 (500 μM), we observed luminescence from the HRP-loaded GUVs (Figure 2e) but not if we used the buffer taken from above the GUVs that were settled before. The photon counts gradually increased over 15 min after the addition of the substrates and remained stable for over 1 h. The reaction profile inside the GUVs was different from that in solution (Figure 2c), where luminescence decayed exponentially with time. The gradual increase and relatively long constant luminescence production inside the GUVs were presumably due to the constant diffusion of the substrates into the GUVs, lower overall HRP concentrations, and permeation dynamics of luminol and H2O2. To access the time scale of substrate transfer across the membrane, we encapsulated a H2O2 sensor inside a GUV and measured its response time after adding H2O2 externally (Figure S3). Indeed, the sensor only responded 10 min after adding H2O2 and saturated at ca. 20 min, which is similar to the time profile of the intracellular chemiluminescence observed in Figure 2e. Building on these results, we functionalized two populations of GUVs (POPC with 10 mol % POPG, 1% DGS-NTA-Ni2+, and 0.1% membrane dye) at their outer membrane with either iLID (membrane labeled with DiL, shown in green) or Nano (membrane labeled with DiD, shown in red) through the binding of His6-tags on the proteins to the DSG-NTA-Ni2+ lipids. Moreover, the iLID-functionalized GUVs were loaded with HRP, such that the chemiluminescence reaction was localized inside them. In an equal mixture of iLID and Nano GUVs (Figure S4), the chemiluminescence reaction was initiated upon addition of H2O2 and luminol; as a result, iLID and Nano GUVs formed heteroclusters (Figure 2f). Also, the 3D view of these GUVs showed membrane deformation and large contact sites between the iLID- and Nano-functionalized GUVs under these conditions (Figure S5). For comparison, iLID and Nano GUVs remained separated in the dark if the substrates were not added and clustered similarly to the chemiluminescence sample if illuminated with blue light externally. In fact, the extent of GUV clustering was as efficient with chemiluminescence as it was with external blue light and was much higher than in the dark, as quantified from the area occupied by the GUV clusters (Figure 2g). The contribution of homoclustering among either iLID or Nano GUVs was low under all conditions and not dependent on the chemiluminescence or the illumination. Some homophilic clustering is inevitable due to GUVs statistically settling down next to each other and GUVs of the same type being pulled together by heterophilic interactions. As a control, we encapsulated HRP inside Nano-functionalized GUVs instead of the iLID-functionalized ones, which would require intercellular photoactivation of iLID to observe GUV clustering. In this case, the GUV clustering was still higher than in the dark but much weaker than in the case of intracellular photoactivation (Figure 2f,g), speaking for the distance dependence of the chemiluminescence photoactivation mechanism. These findings together show that the intracellular chemiluminescence generated inside the sender GUVs is sufficient and is as efficient as direct blue light illumination in mediating heterophilic GUV-GUV adhesions.

H2O2 is an important mediator of redox signaling in living cells and a common signaling molecule used in communities of synthetic cells. In this work, H2O2 is not just used as an intracellularly produced signaling molecule but also as a regulator of adhesion between sender and receiver GUVs that brings the two within signaling range (Figure 3a). This dual use of H2O2 as both a membrane-permeable signaling molecule and a reactant of the chemiluminescence reaction allowed us to spatiotemporally link these two functions. To coregulate signal production and spatial proximity as a function of intracellular enzymatic activity, we generated H2O2 inside the GUVs (Figure 3b). To achieve this, we encapsulated the enzyme glucose oxidase (GOx, 100 mU/ml) inside the GUVs (POPC with 10% POPG and 0.1% DiL) and incorporated α-hemolysin (α-HL, 15 ng/μL) pores to enable the transport of its substrate, glucose. GOx oxidizes glucose to gluconic acids and produces H2O2, where we monitored intracellular H2O2 production with the protein-based sensor roGFP2-Orp1 (20 μM). roGFP2-Orp1 is a ratiometric fluorescent H2O2 sensor, in which the ratio of fluorescence emission upon 405 nm excitation to 488 nm excitation increases as the sensor is oxidized by H2O2 (IEx 405 nm/IEx 488 nm) (Figure S6a,b). This sensor provides a concentration-independent readout for H2O2 dynamics and is retained inside the GUVs despite the α-HL pores due to its large size. Upon addition of glucose (20 mM), the ratiometric signal of roGFP2-Orp1 inside the GUVs increased over the course of 30 min, as monitored with confocal microscopy (Figure 3b,c and Movie 1). The increased error bar with time was attributed to the nonuniform diffusion and transport of glucose into the GUVs within the sample. Subsequently, we converted the H2O2 generated inside the GUVs into a light signal by using the HRP-catalyzed chemiluminescence reaction. To this end, the enzymes HRP (2 μg/mL) and GOx (100 mU/ml) were coencapsulated into the GUVs, and the substrates luminol (200 μM) and glucose (50 mM) were added externally (Figure 3d). Chemiluminescence from GUVs increased over 60 min and subsequently decayed over a couple of hours as measured with a plate reader. Yet, when a higher GOx concentration (500 mU/ml) was encapsulated into the GUV, the chemiluminescence was higher and decayed already in 3 h rather than 4 h. Similarly, the chemiluminescence output from the GUVs could be modulated by changing the concentration of glucose (0–500 mM) or HRP (0–2 μg/mL) (Figure S7). We found that at concentrations above 50 mM glucose and 2 μg/mL HRP the chemiluminescence saturated. Next, we investigated how the adhesions between sender and receiver GUVs correlated with H2O2 production in the sender GUVs and whether the coupled chemiluminescence provides a self-regulatory mechanism for spatial proximity. Sender GUVs (membrane shown in green) were loaded with GOx (100 or 500 mU/mL) and HRP (2 μg/mL), equipped with α-HL pores (15 ng/μL), and functionalized with iLID. Receiver GUVs (membrane shown in red) were functionalized with Nano (Figure 3e and Figure S8). Upon addition of glucose (50 mM) and luminol (200 μM) under mild shaking (30 rpm), the sender and receiver GUVs formed intermixed clusters as a result of the chemiluminescence in the sender GUVs. In this and all subsequent experiments, the mild shaking was essential for the GUVs to move in the suspension and avoid settling down. Control GUVs that were kept in the dark and did not produce chemiluminescence did not cluster with each other. The degree of GUV-GUV clustering increased with time peaking at different times depending on the GOx concentration in the sender GUVs (Figure 3f). While samples with senders that produce more chemiluminescence faster (500 mU/mL GOx) the GUVs clustered more and already fully after 1 h, in samples with senders that produce less and more sustained chemiluminescence (100 mU/mL GOx) the GUV clustering increased over the course of 3 h. Later, as the chemiluminescence output decreased, GUV-GUV clustering also decreased after 4 h in both samples. The temporally dynamic clustering of the GUVs followed the profile of the chemiluminescence reaction in sender GUVs with different concentrations of GOx (Figure 3d); i.e., the GUVs came together as long as the sender GUVs produced chemiluminescence and kept the photoswitchable adhesions activated. Overall, the coupled production of H2O2 and GUV-GUV adhesions allows for adaptive multicellularity and self-regulation of the communication time scale.

Bringing the receiver into the signaling range of the sender and keeping a sustained proximity are important regulators of signaling efficiency. The earlier findings already showed that localized chemiluminescence inside the sender cell reversibly modulates adhesion to a receiver cell as a function of H2O2 production. Next, we wanted to show that the H2O2 from the sender cell can also act as a signaling molecule and that transmission efficiency depends on its self-regulated proximity to the receiver (Figure 4a). To do so, we mixed equal numbers of sender and receiver GUVs under mild shaking (30 rpm) and quantified the response of the receiver to H2O2 production in the sender. The sender GUVs (shown in green) contained GOx (100 mU/mL) for H2O2 production and HRP as the chemiluminescence catalyst and had α-HL pores in their membranes for substrate permeability. In addition, they were functionalized with iLID on their outer surface. The receiver GUVs (shown in red) were loaded with the H2O2 sensor roGFP2-Orp1 and were functionalized with Nano on their outer surface. After we added glucose and luminol, the ratiometric response of the H2O2 sensor increased in the receiver GUVs that were directly attached and sustained close proximity to sender GUVs, i.e. proximal (Figure 4b,c). The response of the H2O2 sensor in the receiver GUVs was distance dependent, and receivers that were not attached to senders, i.e. distal, did not respond to the H2O2 signal efficiently. Moreover, chemiluminescence that triggered adhesion between sender and receiver GUVs was essential for the H2O2 signal to reach the receiver GUVs: if functionalization with iLID and Nano was omitted, the receivers did not respond, even if they were occasionally proximal to the sender cells. In fact, the response in the absence of adhesions was similar to that when the sender did not produce a H2O2 signal for GUVs when GOx was left out. Altered signal transduction efficiency when adhesion occurred was in part due to the transfer of α-HL pores from the sender to the GUV upon adhesion, as detailed below. Herein, it should also be noted that roGFP2-Orp1 has only micromolar sensitivity to H2O2 and takes >5 min to react (Figure S6b). These features minimized nonspecific signaling between GUVs that were transiently in proximity as the GUVs moved in the sample (30 rpm shaking) and contributed to the specific signaling between GUVs with sustained proximity based on the iLID-Nano binding. To access the signaling range from the GUV membrane, we used Ca2+ ions as a model soluble signal and monitored its release upon addition of an ionophore with the calcium-sensitive fluorescent indicator Rhod2 (Figure S9). We observed that the fluorescent signal from the Rhod2-Ca2+ complex inversely correlates with the distance from the GUV membrane and that the signaling was much stronger in close proximity of the membrane (d < 1 μm) than farther away. Overall, these results show how the H2O2 signal that is produced plays a dual role in communication in acting as a direct chemical signal and in regulating the proximity of the sender and receiver to an efficient signaling range.

An important feature of cell-to-cell communication is its bidirectionality, which allows both players to benefit from the exchange and can be an integral part of feedback mechanisms in multicellular communities. That is, the receiver cell replies to the initial sender cell with a secondary signaling molecule. To close the loop of communication, it is necessary to implement a mechanism that releases the secondary signal in response to the primary signal (Figure 5c). We proposed, as a mechanism of adaptation that enables backward signaling, altering the permeability of the receiver GUVs, depending on contact with the sender GUVs that contain α-HL pores. In initial experiments, we mixed sender GUVs (membrane shown in green) that had α-HL pores and were functionalized with iLID with receiver GUVs (membrane shown in cyan) that were loaded with a non-membrane-permeable fluorescent dye, sulfo-Cy5 (shown in red) and were functionalized with Nano (Figure 5a). Under blue light illumination, receiver GUVs that encountered sender GUVs became leaky and released the internal sulfo-Cy5 dye. This permeabilization was clearly contact dependent and was not observed for receiver GUVs that were not in contact with sender GUVs (Figure 5b). Moreover, if the sender GUVs did not have α-HL but still the light triggered adhesions and/or if external H2O2 was added, there was no increase in receiver permeability (Figure 5a,b). These findings exclude increased membrane tension and lipid oxidation as alternative mechanisms that could increase the membrane permeability and show that α-HL is essential for the contact-dependent increase in permeability. Additional proof for contact-dependent permeabilization was provided by experiments in which sulfo-Cy5 was added externally to the aforementioned GUVs, and a sulfo-Cy5 signal inside receiver GUVs that were in contact with sender GUVs was bleached (Figure S10 and Movie 2). The fluorescent recovery after photobleaching in receiver GUVs was observable within minutes, showing that an exchange occurred with the external sulfo-Cy5 pool. At this point, we added bidirectionality to the established self-regulated communication. We chose the Ca2+ ion, as it is another important messenger molecule in cells and serves as a non-membrane-permeable backward signal to be transmitted from the initial receiver GUVs to the sender GUVs. Therefore, Ca2+ (20 μM) was loaded into the receiver GUVs (membrane shown in red, functionalized with Nano, loaded with the H2O2 sensor roGFP2-Orp1, shown in cyan). To detect the Ca2+ signal, we modified the sender GUVs (membrane shown in green, membrane functionalized with iLID and α-HL, loaded with GOx and HRP) with a lipophilic calcium indicator, Calcium Green C24 (shown in purple) (Figure 5c). In preliminary experiments, we verified that the fluorescence of Calcium Green C24 in the GUV membranes increased upon addition of external Ca2+ (Figure S11 and Movie 3). These sender and receiver GUVs were mixed in equal portions, and the production of the primary signal, H2O2, and the coupled chemiluminescence in the sender GUVs were triggered with glucose and luminol. In these communities, sender and receiver GUVs formed heterophilic adhesions as a result of chemiluminescence activation, together with a two-way signal exchange between the different GUV populations (Figure 5c,d). More precisely, the H2O2 produced in the sender GUVs (shown in green) resulted in an increased ratiometric response from the H2O2 sensor, roGFP2-Orp1 (shown in cyan), in the receiver GUVs (shown in red). At the same time, the Ca2+ ions released from the receiver GUVs upon adhesion caused an increase in the fluorescence of the Ca2+ sensor Calcium Green C24 (shown in purple) in the membrane of the sender GUVs. Both H2O2 and Ca2+ signaling were observed only when sender and receiver GUVs were in close proximity, and distal sender or receiver GUVs did not respond (Figure 5e). Notably, both the H2O2 sensor roGFP2-Orp1 and the calcium sensor Calcium Green C24 have the same excitation wavelength at 488 nm, but the different spatial localization of these fluorophores (roGFP2-Orp1 in the lumen of the receiver GUV vs Calcium Green C24 in the membrane of the sender GUV) allowed us to distinguish between the two signals. To verify that there was no crosstalk between the two channels, we repeated the experiment without roGFP2-Orp1 in the lumen of receiver GUVs (Figure S12). In agreement with the previous result, chemiluminescence triggered adhesions between sender and receiver GUVs and resulted in the transfer of the secondary Ca2+ signal to the sender GUVs. To exclude the possibility that the released Ca2+ is responsible for the GUV clustering, we repeated the experiment without including the photoswitchable proteins and releasing Ca2+ from the GUVs with ionomycin (Figure S13) but the GUVs remained well dispersed before and after the release of the Ca2+ ions. Overall, these findings showed how production of the primary signal self-regulates both the spatial relationship and the two-way communication between sender and receiver GUVs.

Here we presented adaptive cell-to-cell communication that is self-regulated and bidirectional. The first layer of regulation in this two-membered GUV community derives from the spatial proximity of sender and receiver being a function of the signal produced in the sender GUV. Sender and receiver therefore adhere only to each other, come into signaling range for the duration of H2O2 signal production, and separate once the signal weakens, yet keep a memory of the exchange. Similar coupling of signal production and adhesion is widely found in biology, the dynamic adhesions of different cells of the adaptive immune system being a prime example. The second layer of regulation occurs as a result of the change in permeability of the receiver GUV once it has adhered to the sender GUV. This allows the receiver GUV to signal backward only when it is in close proximity with the sender and renders the two-way communication efficient. These design principles behind self-regulated proximity and adaptive two-way communication can be implemented in other communities to make communication more effective and dynamic. Our results demonstrate that endogenous chemiluminescence activates the photoswitchable protein iLID. Such proteins are usually activated through external illumination in optogenetic studies, which limits applications to transparent and nonscattering systems. Recently, in situ produced bioluminescent light was discovered as an alternative mode of photoactivation. Here, we demonstrate that the chemiluminescence reaction between luminol and H2O2 is a third way to activate light-responsive proteins. Chemiluminescence has the clear advantage of not needing genetic engineering and being well-established for bioimaging. Therefore, chemiluminescence photoactivation could enable targeting of optogenetic tools to sites of increased redox stress and disease that are inaccessible to light. We envision that the herein demonstrated adaptive communication and chemiluminescence activation of optogenetic systems will give rise to effective multicellular communities, including both synthetic and living cells, and applications in biomedicine, including bioimaging and targeted drug delivery.

POPC (1-palmitoyl-2-oleoyl-glycero-3-phosphocholine, Cat. No. 850457), POPG (1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1′-rac-glycerol) sodium salt, Cat# 840457), DGS-NTA-Ni (1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid)succinyl] nickel salt, Cat# 790404), 18:1 Biotinyl Cap PE (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(cap biotinyl) (sodium salt), Cat. No. 870273) in chloroform were purchased from Avanti Polar Lipids. Amplex Red (Amplex Red reagent, Cat. No. A12222), calcium-sensitive dye Rhod-2 (tripotassium salt, cell impermeant, Cat. No. R14220), streptavidin (Cat No. 434301) and the membrane dyes DiO (3,3′-dioctadecyl-oxacarbocyanine perchlorate, Cat. No. D4292), DiL (1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate, Cat. No. D3911), and DiD (1,1′-dioctadecyl-3,3,3′,3′-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate salt, Cat. No. D7757) were purchased from Thermo-Fisher Scientific. Chemicals and enzymes including luminol (Cat# 123072), H2O2 (Cat. No. 216763), Chelex 100 sodium form (Cat. No. C7901), α-HL (α-hemolysin from Staphylococcus aureus, Cat. No. H9395), HRP (peroxidase from horseradish, Cat. No. P6782), GOx (glucose oxidase from Aspergillus niger, Cat. No. G7141), and ionomycin (ionomycin calcium salt from Streptomyces conglobatus, Cat. No. I0634) were purchased from Sigma-Aldrich. Sulfo-Cy5 (Sulfo-Cyanine5 carboxylic acid, Cat. No. 13390) was purchased from Lumiprobe. Calcium Green C24 (Cat. No. ab275322) was purchased from Abcam. Microscopy experiments with GUVs were performed in 18-well μ-slide glass bottom chambers (Cat. No. 80827) from ibidi.

pQE-80L iLID (C530M) and pQE-80L MBP-SspB Nano were gifts from Brian Kuhlman (Addgene Nos. 60408 and 60409, respectively). pETG10A roGFP2-Orp1 was a gift from Prof. Markus Schwarzländer. All proteins were expressed in E. coli BL21 (DE3) and purified using Ni2+-NTA affinity chromatography as reported in references.

GUVs were prepared using the gel-assisted rehydration method by poly(vinyl alcohol) (PVA) as previously described. For this, 5% (w/v) PVA (MW: 145000 g/mol) was dissolved in Milli-Q water by overnight shaking at 400 rpm at 80 °C. 40 μL of PVA solution was spread on a glass slide (60 mm × 24 mm) and dried at 50 °C for 30 min to form a thin film. Then, 5 μL of a 10 mg/mL lipid mixture dissolved in chloroform was spread onto the PVA film and dried at 30 °C for 1 h. The lipid mixture contained POPC supplemented with 10 mol % POPG, and 0.1 mol % DGS-NTA-Ni2+ if the GUVs were functionalized with the proteins and 0.2 mol % Dil, DiD, or DiO dye. A chamber was assembled on top of the dried film using a self-made Teflon spacer and a second glass slide. The lipid films were rehydrated for 1 h at RT with 1 mL of buffer A (100 mM NaCl, 10 mM Tris, pH 7.4) with 100 mM sucrose or raffinose as well as all the components to be loaded into the GUV depending on the experiment (100 mU/mL GOx, 2 μg/mL HRP, 15 ng/μL α-HL, 20 μM roGFP2-Orp1, 10 μM Calcium Green C24, 20 μM CaCl2). For GUV encapsulated with roGFP2-Orp1, 1 mM DTT was added to minimize autoxidation during GUV preparation according to ref (35). To harvest the GUVs, the chamber was inverted for 5 min and the GUVs were collected into a 1.5 mL LoBind Eppendorf tube. If the GUVs were functionalized with iLID or Nano protein, 1 μM of either His6-tagged iLID or His6-tagged Nano was added to 400 μL of GUVs and incubated in the dark for 1 h. To remove the encapsulated components and excess His-tagged protein from the external solution, 400 μL of GUVs was washed three times by adding 1 mL of washing buffer, spinning the GUVs down (1000 rpm for 5 min), and removing the upper 1 mL (remaining 2% of the initial concentrations). For GUVs with encapsulated GOx, the rehydration buffer contained 100 mM raffinose and buffer A with 67.8 mM KCl was used as a washing buffer in order to balance the osmotic pressure and create a density difference between the GUVs and the outer solution. For the other GUVs without loaded GOx, the rehydration buffer contained 100 mM sucrose and buffer A with 100 mM glucose was used as the washing buffer. For GUVs involving Ca2+ communication, buffer A was pretreated with 50 mg/mL Chelex 100 sodium form to remove any Ca2+ contamination and the washing buffer contained 4 mM EGTA.

The chemiluminescence and fluorescence anisotropy were measured in white- and black-bottom 96-well plates, respectively, using a multimode plate reader (Spark, Tecan Life Science). 200 μL of GUVs loaded with HRP (2 μg/mL) without membrane dye or the external buffer (negative control) was mixed with luminol (200 μM) and H2O2 (500 μM), and the chemiluminescence was measured in the range of 400–500 nm. The competitive fluorescence polarization assay was based on previously published protocols. Briefly, 200 nM SsrA-TAMRA peptide (Sequence: AANDENY-TAMRA) was mixed with 1 μM Nano in buffer (Vtotal = 200 μL) and titrated with increasing concentrations of iLID (0–25 μM). The fluorescence polarization was measured (λEx = 540 nm, λEm = 590 nm) in the dark and after 2 min blue light illumination. Subsequently, the fluorescence polarization measurement was repeated in the presence of 5 μM iLID and the added components of the chemiluminescence reaction (200 μM luminol, 500 μM H2O2, and 2 μg/mL HRP).

The appropriate sender and receiver GUV populations were mixed in an approximately 1:1 ratio in μ-slide 18-well glass-bottom chambers (Vtotal = 60 μL). The chemiluminescence reaction was initiated through the addition of 200 μM luminol and 500 μM H2O2 in GUV populations without encapsulated GOx. For GUV populations with encapsulated GOx, equal volumes of iLID functionalized GUVs with encapsulated GOx (in 67.8 mM KCl in buffer A) and Nano functionalized GUVs (in 100 mM glucose in buffer A) were mixed to achieve a final glucose concentration of 50 mM and 200 μM luminol was added. Samples without the chemiluminescence reaction were kept in the dark or under blue light illumination as negative and positive controls. The samples were incubated on an orbital shaker at 30 rpm for up to 4 h. In different experiments the clustering of the GUVs (DiD and DiL channels), internal GUV fluorescence intensity from the roGFP2-Orp1 sensor (membrane labeled with DiD) or sulfo-Cy5 (membrane labeled with DiO), and fluorescence intensity of the Calcium Green C24 were quantified using confocal microscopy.

All images were acquired on a Leica SP8 confocal laser scanning microscope through a 63× water objective equipped with four laser lines (405, 488, 552, and 638 nm). The following exitation and emmision parameters were used for imaging: roGFP2-Orp1 sensor (λEx = 405 nm or λEx = 488 nm, λEm = 590–630 nm), Calcium Green C24 (λEx = 488 nm, λEm = 510–550 nm), DiO (λEx = 488 nm, λEm = 510–550 nm), DiL dye (λEx = 552 nm, λEm = 560–630 nm), DiD dye (λEx = 638 nm, λEm= 650–700 nm), sulfo-Cy5 (λEx = 638 nm, λEm = 650–700 nm). GUV-GUV clustering was analyzed by converting the images in the DiL and DiD channels into binary images and merging the two channels. Subsequently, the Fill Holes function was used to fill out all GUVs, and the Particle Analyzer tool was used to detect the number and area of objects larger than 10 μm2. All images were analyzed using ImageJ 1.52b.

The data were plotted as the mean ± SEM from technical triplicates. The statistical significance was determined by one-way ANOVA through OriginPro 9.1. p values: ns >0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.