Protein Interactions In Living Cells Research Articles

Erratum: Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission.

An erratum was issued for:Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission. The Authors section was updated from: György Vámosi1 Sarah Miller2 Molika Sinha2 Gabor Mocsár1 Malte Renz2 1Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen 2Gynecologic Oncology Division, Stanford University School of Medicine to: György Vámosi1 Sarah Miller2 Molika Sinha2 Maria Kristha Fernandez2 Gabor Mocsár1 Malte Renz2 1Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen 2Gynecologic Oncology Division, Stanford University School of Medicine.

Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission.

Förster Resonance Energy Transfer (FRET) is the radiationless transfer of energy from an excited donor to an acceptor molecule and depends upon the distance and orientation of the molecules as well as the extent of overlap between the donor emission and acceptor absorption spectra. FRET permits to study the interaction of proteins in the living cell over time and in different subcellular compartments. Different intensity-based algorithms to measure FRET using microscopy have been described in the literature. Here, a protocol and an algorithm are provided to quantify FRET efficiency based on measuring both the sensitized emission of the acceptor and quenching of the donor molecule. The quantification of ratiometric FRET in the living cell not only requires the determination of the crosstalk (spectral spill-over, or bleed-through) of the fluorescent proteins but also the detection efficiency of the microscopic setup. The protocol provided here details how to assess these critical parameters.

Direct measurement of protein-protein interactions by FLIM-FRET at UV laser-induced DNA damage sites in living cells.

Protein–protein interactions are essential to ensure timely and precise recruitment of chromatin remodellers and repair factors to DNA damage sites. Conventional analyses of protein–protein interactions at a population level may mask the complexity of interaction dynamics, highlighting the need for a method that enables quantification of DNA damage-dependent interactions at a single-cell level. To this end, we integrated a pulsed UV laser on a confocal fluorescence lifetime imaging (FLIM) microscope to induce localized DNA damage. To quantify protein–protein interactions in live cells, we measured Förster resonance energy transfer (FRET) between mEGFP- and mCherry-tagged proteins, based on the fluorescence lifetime reduction of the mEGFP donor protein. The UV-FLIM-FRET system offers a unique combination of real-time and single-cell quantification of DNA damage-dependent interactions, and can distinguish between direct protein–protein interactions, as opposed to those mediated by chromatin proximity. Using the UV-FLIM-FRET system, we show the dynamic changes in the interaction between poly(ADP-ribose) polymerase 1, amplified in liver cancer 1, X-ray repair cross-complementing protein 1 and tripartite motif containing 33 after DNA damage. This new set-up complements the toolset for studying DNA damage response by providing single-cell quantitative and dynamic information about protein–protein interactions at DNA damage sites.

Exploring GPCR-arrestin interfaces with genetically encoded crosslinkers.

β‐arrestins (βarr1 and βarr2) are ubiquitous regulators of G protein‐coupled receptor (GPCR) signaling. Available data suggest that β‐arrestins dock to different receptors in different ways. However, the structural characterization of GPCR‐arrestin complexes is challenging and alternative approaches to study GPCR‐arrestin complexes are needed. Here, starting from the finger loop as a major site for the interaction of arrestins with GPCRs, we genetically incorporate non‐canonical amino acids for photo‐ and chemical crosslinking into βarr1 and βarr2 and explore binding topologies to GPCRs forming either stable or transient complexes with arrestins: the vasopressin receptor 2 (rhodopsin‐like), the corticotropin‐releasing factor receptor 1, and the parathyroid hormone receptor 1 (both secretin‐like). We show that each receptor leaves a unique footprint on arrestins, whereas the two β‐arrestins yield quite similar crosslinking patterns. Furthermore, we show that the method allows defining the orientation of arrestin with respect to the GPCR. Finally, we provide direct evidence for the formation of arrestin oligomers in the cell.

NMR Characterization of Surface Receptor Protein Interactions in Live Cells Using Methylcellulose Hydrogels.

Interactions of transmembrane receptors with their extracellular ligands are essential for cellular communication and signaling and are therefore a major focus in drug discovery programs. The transition from in vitro to live cell interaction studies, however, is typically a bottleneck in many drug discovery projects due to the challenge of obtaining atomic‐resolution information under near‐physiological conditions. Although NMR spectroscopy is ideally suited to overcome this limitation, several experimental impairments are still present. Herein, we propose the use of methylcellulose hydrogels to study extracellular proteins and their interactions with plasma membrane receptors. This approach reduces cell sedimentation, prevents the internalization of membrane receptors, and increases cell survival, while retaining the free tumbling of extracellular proteins.

NMR Characterization of Surface Receptor Protein Interactions in Live Cells Using Methylcellulose Hydrogels

AbstractInteractions of transmembrane receptors with their extracellular ligands are essential for cellular communication and signaling and are therefore a major focus in drug discovery programs. The transition from in vitro to live cell interaction studies, however, is typically a bottleneck in many drug discovery projects due to the challenge of obtaining atomic‐resolution information under near‐physiological conditions. Although NMR spectroscopy is ideally suited to overcome this limitation, several experimental impairments are still present. Herein, we propose the use of methylcellulose hydrogels to study extracellular proteins and their interactions with plasma membrane receptors. This approach reduces cell sedimentation, prevents the internalization of membrane receptors, and increases cell survival, while retaining the free tumbling of extracellular proteins.

Proximity labeling to detect RNA-protein interactions in live cells.

RNA biology is orchestrated by the dynamic interactions of RNAs and RNA‐binding proteins (RBPs). In the present study, we describe a new method of proximity‐dependent protein labeling to detect RNA–protein interactions [RNA‐bound protein proximity labeling (RBPL)]. We selected the well‐studied RNA‐binding protein PUF to examine the current proximity labeling enzymes birA* and APEX2. A new version of birA*, BASU, was used to validate that the PUF protein binds its RNA motif. We further optimized the RBPL labeling system using an inducible expression system. The RBPL (λN‐BASU) labeling experiments exhibited high signal‐to‐noise ratios. We subsequently determined that RBPL (λN‐BASU) is more suitable than RBPL (λN‐APEX2) for the detection of RNA–protein interactions in live cells. Interestingly, our results also reveal that proximity labeling is probably capable of biotinylating proximate nascent peptide.

Measuring Small-molecule Inhibition of Protein Interactions in Live Cells Using FLIM-FRET.

This protocol was designed to quantitatively measure small-molecule displacement of proteins in live mammalian cells using fluorescence lifetime imaging microscopy-Förster resonance energy transfer (FLIM-FRET). Tumour cell survival is often dependent on anti-apoptotic proteins, which bind to and inhibit pro-apoptotic proteins, thus preventing apoptosis. Small-molecule inhibitors that selectively target these proteins (termed BH3-mimetics) are therefore a promising avenue for the treatment of several cancers. Previous techniques used to study the efficacy of these drugs often use truncated versions of both pro- and anti-apoptotic proteins, as they are membrane bound and hydrophobic in nature. As a result, the true efficacy of these drugs to displace full-length pro-apoptotic proteins in their native environment within a cell is poorly understood. This protocol describes FLIM-FRET methods to directly measure the displacement (or lack of displacement) of full-length Bcl-2 family proteins in live mammalian cells.

Optimizing the fragment complementation of APEX2 for detection of specific protein-protein interactions in live cells

Dynamic protein-protein interactions (PPIs) play crucial roles in cell physiological processes. The protein-fragment complementation (PFC) assay has been developed as a powerful approach for the detection of PPIs, but its potential for identifying protein interacting regions is not optimized. Recently, an ascorbate peroxidase (APEX2)-based proximity-tagging method combined with mass spectrometry was developed to identify potential protein interactions in live cells. In this study, we tested whether APEX2 could be employed for PFC. By screening split APEX2 pairs attached to FK506-binding protein 12 (FKBP) and the FKBP12-rapamycin binding (FRB) domain, which interact with each other only in the presence of rapamycin, we successfully obtained an optimized pair for visualizing the interaction between FRB and FKBP12 with high specificity and sensitivity in live cells. The robustness of this APEX2 pair was confirmed by its application toward detecting the STIM1 and Orial1 homodimers in HEK-293 cells. With a subsequent mass spectrometry analysis, we obtained five different biotinylated sites that were localized to the known interaction region on STIM1 and were only detected when the homodimer formed. These results suggest that our PFC pair of APEX2 provides a potential tool for detecting PPIs and identifying binding regions with high specificity in live cells.

Weak protein–protein interactions in live cells are quantified by cell-volume modulation

Weakly bound protein complexes play a crucial role in metabolic, regulatory, and signaling pathways, due in part to the high tunability of their bound and unbound populations. This tunability makes weak binding (micromolar to millimolar dissociation constants) difficult to quantify under biologically relevant conditions. Here, we use rapid perturbation of cell volume to modulate the concentration of weakly bound protein complexes, allowing us to detect their dissociation constant and stoichiometry directly inside the cell. We control cell volume by modulating media osmotic pressure and observe the resulting complex association and dissociation by FRET microscopy. We quantitatively examine the interaction between GAPDH and PGK, two sequential enzymes in the glycolysis catalytic cycle. GAPDH and PGK have been shown to interact weakly, but the interaction has not been quantified in vivo. A quantitative model fits our experimental results with log Kd = -9.7 ± 0.3 and a 2:1 prevalent stoichiometry of the GAPDH:PGK complex. Cellular volume perturbation is a widely applicable tool to detect transient protein interactions and other biomolecular interactions in situ. Our results also suggest that cells could use volume change (e.g., as occurs upon entry to mitosis) to regulate function by altering biomolecular complex concentrations.

Real time imaging of tri‐molecular protein interactions in live cells by Förster resonance energy transfer (FRET) microscopy

Malignant cancer cells frequently secrete significant amounts of transforming growth factor beta (TGF‐beta), hyaluronan (HA) and hyaluronidases to facilitate metastasizing to target organs. In a non‐canonical signaling, TGF‐beta binds membrane hyaluronidase Hyal‐2 for recruiting tumor suppressors WWOX and Smad4, and the resulting Hyal‐2/WWOX/Smad4 complex is accumulated in the nucleus to enhance SMAD‐promoter dependent transcriptional activity. Yeast two‐hybrid analysis showed that WWOX acts as a bridge to bind both Hyal‐2 and Smad4. When WWOX‐expressing cells were stimulated with high molecular weight HA, an increased formation of endogenous Hyal‐2/WWOX/Smad4 complex occurred rapidly, followed by relocating to the nuclei in 20–40 min. In WWOX‐deficient cells, HA failed to induce Smads relocation to the nucleus. To prove the signaling event, we designed a real time tri‐molecular FRET analysis and revealed that HA induces the signaling pathway from ectopic Smad4 to WWOX and finally to p53, as well as from Smad4 to Hyal‐2 and then to WWOX. An increased binding of the Smad4/Hyal‐2/WWOX complex occurs with time in the nucleus that leads to bubbling cell death. In contrast, HA increases the binding of Smad4/WWOX/p53, which causes membrane blebbing but without cell death. In traumatic brain injury‐induced neuronal death, the Hyal‐2/WWOX complex was accumulated in the apoptotic nuclei of neurons in the rat brains in 24 hr post injury, as determined by immunoelectron microscopy. Together, HA activates the Hyal‐2/WWOX/Smad4 signaling and causes bubbling cell death when the signaling complex is overexpressed.Support or Funding InformationMinistry of Science and Technology, Taiwan

Priming GPCR Signaling Through the Synergistic Effect of Two G Proteins

While individual GPCRs are known to activate one or more G proteins, the GPCR‐G protein interaction is viewed as a bi‐molecular event involving the formation of a ternary ligand‐GPCR‐G protein complex. We have designed a FRET‐based sensor to monitor and modulate GPCR – G protein interactions in live cells. Using this systematic protein affinity strength modulation (SPASM) approach we present evidence that individual GPCR‐G protein interactions can reinforce each other to enhance signaling through canonical downstream second messengers, a phenomenon we term GPCR priming. Specifically, we find that the presence of non‐cognate Gq protein enhances cAMP stimulated by two Gs coupled receptors, β2‐adrenergic receptor (β2‐AR) and D1 dopamine receptor (D1‐R). The C‐terminus of either the Gαs or Gαq subunit is sufficient to enhance Gα subunit activation and cAMP levels. Interaction of Gαs or Gαq C‐termini with the GPCR increases signaling potency suggesting an altered GPCR conformation as the underlying basis for GPCR priming. We propose three parallel mechanisms involving (1) sequential G protein interactions at the orthosteric site, (2) simultaneous allosteric and orthosteric G protein interactions, and (3) asymmetric GPCR dimers. GPCR priming suggests another layer of regulation in the classic GPCR ternary complex model, with broad implications for the multiplicity inherent in signaling networks.Support or Funding InformationResearch was funded by the American Heart Association Scientist Development Grant (13SDG14270009) & the NIH (1DP2 CA186752‐01 & 1‐R01‐GM‐105646‐01‐A1) to S.S. T.M.G is an NCBS‐inStem Career Development Fellow. R.F.S is a Life Sciences Research Foundation postdoctoral fellow.

Protein Micropatterning Assay: Quantitative Analysis of Protein-Protein Interactions.

Characterization, especially quantification, of protein interactions in live cells is usually not an easy endeavor. Here, we describe a straightforward method to identify and quantify the interaction of a membrane protein ("bait") and a fluorescently labeled interaction partner ("prey") (membrane-bound or cytosolic) in live cells using Total Internal Reflection Fluorescence microscopy. The bait protein is immobilized within patterns in the plasma membrane (e.g., via an antibody); the bait-prey interaction strength can be quantified by determining the prey bulk fluorescence intensity with respect to the bait patterns. This method is particularly suitable also for the analysis of weak, transient interactions that are not easily accessible with other methods.

Enabling systematic interrogation of protein-protein interactions in live cells with a versatile ultra-high-throughput biosensor platform.

Large-scale genomics studies have generated vast resources for in-depth understanding of vital biological and pathological processes. A rising challenge is to leverage such enormous information to rapidly decipher the intricate protein-protein interactions (PPIs) for functional characterization and therapeutic interventions. While a number of powerful technologies have been employed to detect PPIs, a singular PPI biosensor platform with both high sensitivity and robustness in a mammalian cell environment remains to be established. Here we describe the development and integration of a highly sensitive NanoLuc luciferase-based bioluminescence resonance energy transfer technology, termed BRET(n), which enables ultra-high-throughput (uHTS) PPI detection in live cells with streamlined co-expression of biosensors in a miniaturized format. We further demonstrate the application of BRET(n) in uHTS format in chemical biology research, including the discovery of chemical probes that disrupt PRAS40 dimerization and pathway connectivity profiling among core members of the Hippo signaling pathway. Such hippo pathway profiling not only confirmed previously reported PPIs, but also revealed two novel interactions, suggesting new mechanisms for regulation of Hippo signaling. Our BRET(n) biosensor platform with uHTS capability is expected to accelerate systematic PPI network mapping and PPI modulator-based drug discovery.

The use of FLIM-FRET for the detection of mitochondria-associated protein interactions.

Fluorescence lifetime imaging microscopy-Förster resonant energy transfer (FLIM-FRET) is a high-resolution technique for the detection of protein interactions in live cells. As the cost of this technology becomes more competitive and methods are devised to extract more information from the FLIM images, this technique will be increasingly useful for studying protein interactions in live cells. Here we demonstrate the use of the ISS-Alba FLIM/FCS confocal microscope, which was custom-built for supervised automation of FLIM data acquisition. We provide a detailed protocol for collecting and analyzing good FLIM-FRET data. As an example, we use FLIM-FRET to detect the interaction between BclXL and Bad at the mitochondrial outer membrane in live MCF7 breast cancer cells.

Downstream Targets of Dopamine Receptor D3 Activation in the Mouse Pancreatic β-Cells

We have shown that dopamine modulates a negative feedback control of insulin secretion from mouse islets by activating the dopamine receptor D3 (DRD3). Receptor activation causes a dose-dependent reduction of the frequency of glucose-triggered intracellular calcium oscillations, which correlates with the observed decrease of insulin secretion. Although it is known that DRD3 can inhibit adenylate cyclase (AC), activate potassium channels, and inhibit calcium channels, the downstream molecular targets of the DRD3 signal transduction cascade in the islet are still unknown. The goal of this study is to identify the main signaling pathway mediating the DRD3-driven inhibition of insulin secretion. Preliminary data suggests that DRD3 signaling in β-cells proceeds through modulation of ion channels rather than by inhibition of AC.We used patch clamp to study the effect of dopamine on the ion channels. Based on these experiments we selected molecular targets to investigate further. We have tagged fluorescent proteins on to the Gβγ complex of the heterotrimeric G protein, and also on to candidate-target ion channels. This allows us to use quantitative fluorescence analysis tools to measure the changes in protein-protein interaction before and after pharmacological stimulation of DRD3. Data is analyzed using a spatial intensity distribution analysis (Godin, A. G. et al., PNAS, 2011) to detect protein interactions in live cells and fixed samples. This analysis allow us to compare results from cell lines overexpressing our proteins of interest with the results from native pancreatic islets from mice and human donors where immuno-staining techniques can be used to label the proteins under investigation.The information gained will help us understand how pancreatic islets regulate insulin secretion. Moreover, this signaling pathway could potentially be exploited to design effectors of insulin secretion as a treatment for type-2 diabetes.

Investigating γ-secretase protein interactions in live cells using active site-directed clickable dual-photoaffinity probes

Clickable γ-secretase active site-directed dual-photoaffinity probes specifically label components of the γ-secretase complex and form crosslinks between PS1-NTF and PS1-CTF.

Bioluminescence Assisted Switching and Fluorescence Imaging (BASFI)

Förster resonance energy transfer (FRET) and bioluminescence resonance energy transfer (BRET) are two major biophysical techniques for studying nanometer-scale motion dynamics within living cells. Both techniques read photoemission from the transient RET-excited acceptor, which makes RET and detection processes inseparable. We here report a novel hybrid strategy, bioluminescence assisted switching and fluorescence imaging (BASFI) using a bioluminescent Renilla luciferase RLuc8 as the donor and a photochromic fluorescent protein Dronpa as the acceptor. When in close proximity, RET from RLuc8 switches Dronpa from its original dark state to a stable bright state, whose fluorescence is imaged subsequently with an external laser. Such decoupling between RET and imaging processes in BASFI promises high photon flux as in FRET and minimal bleedthroughs as in BRET. We demonstrated BASFI with Dronpa-RLuc8 fusion constructs and drug-inducible intermolecular FKBP-FRB protein–protein interactions in live cells with high sensitivity, resolution, and specificity. Integrating the advantages of FRET and BRET, BASFI will be a valuable tool for various biophysical studies.

Tracking Image Correlation: Combining Single-Particle Tracking and Image Correlation

The interactions and coordination of biomolecules are crucial for most cellular functions. The observation of protein interactions in live cells may provide a better understanding of the underlying mechanisms. After fluorescent labeling of the interacting partners and live-cell microscopy, the colocalization is generally analyzed by quantitative global methods. Recent studies have addressed questions regarding the individual colocalization of moving biomolecules, usually by using single-particle tracking (SPT) and comparing the fluorescent intensities in both color channels. Here, we introduce a new method that combines SPT and correlation methods to obtain a dynamical 3D colocalization analysis along single trajectories of dual-colored particles. After 3D tracking, the colocalization is computed at each particle’s position via the local 3D image cross correlation of the two detection channels. For every particle analyzed, the output consists of the 3D trajectory, the time-resolved 3D colocalization information, and the fluorescence intensity in both channels. In addition, the cross-correlation analysis shows the 3D relative movement of the two fluorescent labels with an accuracy of 30 nm. We apply this method to the tracking of viral fusion events in live cells and demonstrate its capacity to obtain the time-resolved colocalization status of single particles in dense and noisy environments.

Dimerization-Dependent Green and Yellow Fluorescent Proteins

Dimerization-dependent fluorescent proteins (ddFP) are a recently introduced class of genetically encoded reporters that can be used for the detection of protein interactions in live cells. The progenitor of this class of tools was a red fluorescent ddFP (ddRFP) derived from a homodimeric variant of Discosoma red fluorescent protein. Here, we describe the engineering and application of an expanded palette of ddFPs, which includes green (ddGFP) and yellow (ddYFP) variants. These optimized variants offer several advantages relative to ddRFP including increased in vitro contrast and brightness for ddGFP and increased brightness and a lowered pK a for ddYFP. We demonstrate that both variants are useful as biosensors for protease activity in live cells. Using the ddGFP tool, we generated a highly effective indicator of endomembrane proximity that can be used to image the mitochondria-associated membrane (MAM) interface of endoplasmic reticulum (ER) and mitochondria.