Bimolecular Fluorescence Complementation (BiFC) for Live-Cell Imaging of Protein-Protein Interactions

Introduction to BiFC

Protein–protein interactions (PPIs) govern virtually all cellular processes, including signal transduction, gene regulation, and structural assembly. Deciphering the spatial and temporal dynamics of these interactions in living cells is critical for understanding biological function and disease mechanisms. Among the arsenal of tools developed for this purpose, bimolecular fluorescence complementation (BiFC) has emerged as a robust and widely applied approach for visualizing PPIs in real time within intact cells.

BiFC is based on the principle that a fluorescent protein (FP) can be split into two non-fluorescent fragments, each fused to a protein of interest. Upon interaction of these proteins, the two FP fragments reassemble into a functional fluorophore, yielding a fluorescence signal that spatially and temporally reports on the interaction event. BiFC is genetically encoded, does not require external substrates, and can be performed in live cells under physiological conditions.

Molecular Basis of BiFC

Mechanism of Split-Fluorophore Reconstitution

Fluorescent tags like GFP, YFP and mCherry can be split into two parts—an N-terminal piece and a C-terminal piece—that on their own are unstable and won't glow. When each fragment is fused to a pair of interacting proteins and both are made in the same cell, the protein–protein interaction brings the fragments together. This lets them reassemble into the full β-barrel shape, fold correctly, mature their light-emitting core, and regain fluorescence.

The chromophore, formed through autocatalytic cyclization of specific amino acids (e.g., Ser-Tyr-Gly in GFP), requires proper alignment of the FP backbone. Only upon successful reconstitution does the fluorophore become fluorescent. Therefore, fluorescence serves as a direct, irreversible readout of the interaction event.

Kinetics and Thermodynamics of Fragment Association

The kinetics of FP fragment association are slower than typical protein-protein binding due to the requirement for conformational folding and chromophore maturation, which may take several minutes to hours. The process is often driven by the binding affinity of the two proteins of interest rather than the inherent affinity of the FP fragments themselves.

Thermodynamically, complementation is energetically favorable once fragments are in close proximity. This low-energy folding pathway results in a stable fluorophore that resists thermal fluctuations and proteolysis. However, spontaneous self-association of FP fragments—particularly at high expression levels—may occur, underscoring the importance of stringent controls.

Irreversibility and Its Impact on Dynamic Measurements

A defining property of BiFC is the irreversible nature of fluorescence complementation. Once the fluorophore is formed, the structure is highly stable and does not disassemble even if the original interacting proteins separate. This characteristic enhances signal intensity and allows long-term imaging without signal decay.

However, irreversibility introduces a limitation: it prevents real-time observation of interaction dynamics, such as transient association and dissociation cycles. As a result, BiFC is not ideal for capturing rapid, reversible interactions. Instead, it is best suited for detecting stable or cumulative interaction events and mapping subcellular interaction sites.

Engineering Split Fluorescent Protein Pairs

The success of BiFC relies on the effective engineering of split FP pairs that complement with high efficiency, minimal background, and strong signal output. This section discusses key parameters in designing optimal BiFC systems, including scaffold selection, fragment architecture, and protein engineering strategies.

Selection of Fluorophore Scaffolds (GFP, YFP, mCherry, etc.)

Fluorescent proteins used in BiFC must exhibit robust folding, high brightness, and predictable photophysical properties. GFP and its derivatives—yellow (YFP), cyan (CFP), and red (mCherry, mRFP)—are the most commonly used scaffolds due to their well-characterized structures and performance in mammalian cells.

• GFP and YFP: Favored for their rapid maturation and bright fluorescence. YFP is often preferred in dual-color BiFC due to its spectral separation from CFP or mCherry.
• mCherry and mOrange: Red-shifted variants offer reduced autofluorescence and deeper tissue penetration. Their use is suitable for multiplexed detection of multiple PPIs.
• Split sites: Typical split points are between β-strands 7 and 8 or 10 and 11. These allow independent folding of fragments while preserving the potential for complementation.

Optimization of Fragment Length and Folding Efficiency

Effective complementation depends on the structural stability of individual fragments and their ability to reconstitute upon proximity. Split fragments must be:

• Individually non-fluorescent to avoid background signal.
• Stable in the cytosol, resisting aggregation or degradation.
• Optimally sized: Fragments typically range from 100–150 amino acids. Fragment boundaries are selected to avoid disrupting the β-barrel core.

Directed Mutagenesis to Enhance Brightness and Complementation

Engineering efforts have focused on improving the functional properties of split FPs through mutagenesis:

• Enhanced complementation efficiency: Mutations can improve interface compatibility between fragments.
• Increased brightness: Mutations around the chromophore cavity can enhance quantum yield.
• Reduced spontaneous reassembly: Interface mutations reduce background fluorescence from non-specific complementation.
• Faster maturation: Optimized residues near the chromophore accelerate chromophore formation post-complementation.

Figure 1. Schematic representation of the split-YFP /BiFC method based on the YFP (Horstman A, et al., 2014).

Design and Construction of BiFC Expression Vectors

Effective BiFC analysis begins with the careful design of expression vectors to ensure that both fusion partners are produced at appropriate levels, maintain native function, and allow efficient fluorophore reconstitution.

Fusion Tag Positioning

• Tag Orientation: Test both N- and C-terminal fusions of each protein with the FP fragment. One orientation may interfere less with native folding, binding interfaces, or subcellular targeting.
• Reciprocal Constructs: Construct reciprocal pairs (Protein A–N-fragment + Protein B–C-fragment and vice versa) to confirm interaction specificity and rule out steric artifacts.

Linker Design

• Flexible Linkers: Use short Gly-Ser repeats of 8-15 amino acids to separate the FP fragment from the protein domain.
• Length Optimization: Empirically evaluate linker lengths. Too short may sterically hinder complementation; too long may increase background by permitting spontaneous fragment encounters.
• Rigid vs. Flexible Balance: For interaction domains that require precise orientation, consider semi-rigid linkers to restrain movement and improve complementation efficiency.

Promoter and Co-Expression Strategy

Balanced co-expression of both BiFC fusion constructs is essential for efficient fluorophore reconstitution. Strong ubiquitous promoters such as CMV or EF1α are commonly used in transient expression systems. For controlled or tissue-specific expression, inducible promoters or cell-type-specific promoters may be employed. Co-expression strategies include:

• Dual-plasmid transfection, allowing independent expression control but risking expression imbalance.
• Single-plasmid bicistronic vectors using internal ribosome entry sites (IRES) or 2A self-cleaving peptides to ensure stoichiometric expression from a single transcript.

Validation of Expression Constructs

• Western Blot Analysis: Confirm expression and correct molecular weight of each fusion protein using fragment-specific antibodies.
• Fluorescence Controls: Express each FP fragment alone to verify absence of background fluorescence. Co-express complementary fragments fused to known non-interacting proteins as negative controls.
• Functional Assays: Where possible, validate that fusion proteins retain biological activity (e.g., enzymatic assays, localization studies) to ensure that tagging has not disrupted function.

Cell Culture, Transfection, and Expression Optimization

Selection of Appropriate Mammalian Cell Lines

The choice of cell line significantly influences BiFC performance. Ideal host cells should combine high transfection efficiency, well-characterized morphology, and relevance to the biological context of the proteins under investigation. Commonly used cell lines include:

• HEK293T: High transfection efficiency; suitable for overexpression and screening studies.
• HeLa: Robust growth and nuclear imaging capability; commonly used for signaling studies.
• COS-7 and NIH-3T3: Fibroblast-like morphology; useful for cytoskeletal and membrane-associated interactions.
• U2OS: Flat morphology ideal for high-resolution imaging; supports consistent expression levels.

Transient vs. Stable Transfection Protocols

Two primary strategies are used for introducing BiFC constructs into mammalian cells:

• Transient Transfection: Suitable for short-term studies and rapid screening of interaction pairs. Techniques such as calcium phosphate precipitation, lipofection, or electroporation are commonly used.
• Stable Transfection: Preferred for long-term studies, consistent expression, and reduced inter-experimental variability. This involves genomic integration of BiFC constructs using selection markers (e.g., neomycin, puromycin). Lentiviral or retroviral systems are often used to enhance integration efficiency.

Live-Cell Imaging Setup for BiFC

Microscope Configuration and Filter Sets

Fluorescence microscopy is the primary tool for BiFC imaging. Epifluorescence, confocal, or spinning-disk confocal systems are commonly used. Filter sets must match the spectral properties of the reconstituted fluorophore. For example, GFP-based BiFC requires excitation at 488 nm and emission detection near 510 nm. Dual or triple-channel configurations enable co-localization or multi-color BiFC studies.

Environmental Control: Temperature, CO₂, and Humidity

To maintain cell viability and physiological relevance, imaging should be conducted under controlled environmental conditions. Live-cell chambers with 37°C temperature, 5% CO₂, and high humidity are standard. Prolonged imaging sessions necessitate stable environmental control to prevent cell stress and artifactual changes in protein behavior.

Minimizing Phototoxicity and Photobleaching

Repeated excitation of fluorescent proteins may induce phototoxic effects and photobleaching, compromising signal quality. To minimize these effects, low-intensity illumination, short exposure times, and sensitive detectors are recommended. Time-lapse imaging should be optimized to balance temporal resolution with cellular health.

Comparison with FRET, Co-Localization

Applications of BiFC Assay

Mapping Signal Transduction Complexes in Live Cells

BiFC has been instrumental in delineating signaling cascades such as MAPK, Wnt, and TGF-β pathways. The ability to visualize where and when specific interactions occur provides insights into pathway activation, scaffold assembly, and spatial regulation of signaling nodes.

Visualizing Transient Interactions during Cell Cycle Progression

The cell cycle is regulated by a network of transient protein interactions. BiFC enables the capture of such interactions at defined stages, including cyclin-CDK complexes, checkpoint regulators, and chromatin modifiers. These studies have revealed dynamic recruitment patterns and cell cycle-dependent changes in PPI networks.

Studying Disease-Relevant PPIs in Cancer and Neurodegeneration Models

Aberrant PPIs are hallmarks of various diseases. In cancer, BiFC has been applied to study interactions involving oncogenes and tumor suppressors. In neurodegeneration, BiFC has illuminated the oligomerization of proteins such as tau, α-synuclein, and huntingtin in living neurons, supporting drug discovery efforts and mechanistic studies.

Case study

A bimolecular fluorescence complementation flow cytometry screen for membrane protein interactions

Journal: Scientific Reports

Published: 2021

DOI: 10.1038/s41598-021-98810-2

Background

Membrane proteins orchestrate critical cellular processes such as signal transduction, molecular trafficking, and homeostatic regulation. Conventional PPIs assays frequently fail in membranous environments. BiFC overcomes this limitation by reconstituting a split fluorescent protein upon interaction of tagged partners directly within living cells.

Purpose

The authors sought to integrate BiFC with flow cytometry to create a scalable platform capable of detecting and quantifying membrane PPIs in Saccharomyces cerevisiae. This methodology aimed to combine the spatial resolution of BiFC with the throughput and quantitation of cytometric analysis.

Method

• Construct Design: Human aquaporin-0 (hAQP0) and calmodulin (CaM) were each fused to complementary fragments of YFP.
• Yeast Transformation: Fusion constructs were co-transformed into S. cerevisiae and selected on appropriate media.
• Flow Cytometric Analysis: Cells were cultured under optimized conditions, then analyzed by flow cytometry to measure fluorescence complementation.

Results

• Specific Signal Detection: Only yeast co-expressing both hAQP0-YFP_N and CaM-YFP_C exhibited a distinct fluorescent population above autofluorescence levels.
• Background Reduction: Optimization of culture parameters and cytometer gating minimized non-specific signals.
• Quantitative Throughput: The assay distinguished positive from negative interactions in a semi-high-throughput manner without resorting to labor-intensive microscopy.

Figure 2. Bright field and fluorescence images of subcellular localisation of BiFC complexes produced in S. cerevisiae cells.

Figure 3. Flow cytometry quantification of BiFC yields.

Conclusion

The BiFC-flow cytometry workflow furnishes a robust, quantitative platform for mapping membrane PPIs in live yeast cells. Its compatibility with fluorescence-activated cell sorting (FACS) paves the way for library-scale screens to uncover novel interaction partners under physiological conditions.

People Also Ask

What are the main advantages of BiFC?

BiFC offers direct visualization of PPIs without external substrates. It provides high sensitivity, since even weak or low-abundance interactions generate stable fluorescence. It is genetically encoded and compatible with standard fluorescence microscopy

Which fluorescent protein variants are best suited for BiFC?

GFP and YFP variants are commonly used due to efficient folding and brightness. Red‐shifted variants (e.g., mCherry), superfolder GFP (sfGFP), and near‐infrared splits provide advantages in brightness, spectral multiplexing, and imaging in dense tissues

How is BiFC signal intensity quantified?

Signal intensity is measured using fluorescence microscopy coupled with image analysis software. Quantification involves background subtraction, normalization to transfection controls, and statistical evaluation across replicates. Flow cytometry can also be employed in high-throughput formats.

How does linker length affect BiFC efficiency?

Linkers influence the spatial flexibility of fusion proteins. Short linkers may restrict fragment complementation, while long linkers may promote non-specific reconstitution. A typical linker length is 5–20 amino acids, often composed of glycine-serine repeats.

References

• Horstman A, et al. A cautionary note on the use of split-YFP/BiFC in plant protein-protein interaction studies. International journal of molecular sciences, 2014, 15(6): 9628-9643.
• Miller K E, et al. Bimolecular fluorescence complementation (BiFC) analysis: advances and recent applications for genome-wide interaction studies.Journal of molecular biology, 2015, 427(11): 2039-2055.
• Jia Y, et al. Bimolecular Fluorescence Complementation (BiFC) and Multiplexed Imaging of Protein–Protein Interactions in Human Living Cells. Multiplexed Imaging: Methods and Protocols, 2021: 173-190.