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Proximity labeling

Proximity labeling is a powerful biochemical technique for mapping protein-protein interactions and biomolecular neighborhoods in living cells by covalently tagging molecules within nanometer-scale proximity to a genetically targeted protein or subcellular structure. The method involves fusing an engineered enzyme—such as a promiscuous biotin ligase or peroxidase—to a bait protein of interest, which, upon addition of a reactive substrate like biotin or biotin-phenol, labels nearby biomolecules for subsequent affinity purification and identification, often by mass spectrometry or sequencing.^[1] This approach excels at capturing transient, weak, or context-dependent associations that evade traditional immunoprecipitation methods, providing spatial and temporal insights into cellular organization under native conditions.^[2] The foundational proximity labeling strategies emerged in the early 2010s, beginning with BioID in 2012, which utilizes a mutant Escherichia coli biotin ligase (BirA*) to biotinylate lysine residues within approximately 10 nm of the bait over 16–24 hours.^[1] Shortly thereafter, in 2013, APEX was introduced, employing an engineered ascorbate peroxidase to generate short-lived radicals from biotin-phenol in the presence of hydrogen peroxide, enabling rapid labeling (within 1 minute) in a ~20 nm radius but requiring careful control to mitigate oxidative stress. Subsequent advancements, such as TurboID and miniTurboID in 2018, optimized BirA variants through directed evolution for faster (10–60 minutes) and more efficient labeling with lower biotin concentrations, enhancing sensitivity for low-abundance targets while reducing background noise.^[3] Other variants, including peroxidase-based APEX2 and pupylation-based PUP-IT, further expanded the toolkit by improving enzyme size, specificity, and compatibility with diverse substrates like RNA or DNA. Proximity labeling has broad applications across biology, from elucidating mammalian signaling complexes (e.g., nuclear pores or stress granules) to profiling plant immune responses and microbial pathogen-host interfaces, including SARS-CoV-2 interactions.^[2] Its key advantages include in vivo applicability without cell lysis, high spatiotemporal resolution, and the ability to probe insoluble or dynamic structures, though challenges like labeling radius variability, potential artifacts from enzyme promiscuity, and organism-specific optimizations persist.^[4] Recent innovations, such as split-enzyme systems and photocatalytic methods (including energy-transfer photoproximity labeling as of 2025), continue to refine its precision for super-resolution mapping and multi-omics integration.^[4]^[5]

Introduction

Definition and Core Principles

Proximity labeling (PL) is a biochemical technique that employs engineered enzymes or catalysts, genetically fused to or targeted near a protein of interest (often called the "bait"), to covalently tag nearby biomolecules—such as proteins or RNA—within a limited spatial radius, typically using affinity labels like biotin. This method enables the identification of molecular neighbors in living cells without disrupting native structures or interactions, addressing challenges posed by transient or weak associations that are difficult to capture with traditional approaches. By generating reactive species that diffuse only short distances, PL maps local interactomes with high spatiotemporal precision, facilitating insights into cellular organization and dynamics.^[2]^[6] At its core, proximity labeling relies on the localized production of short-lived reactive intermediates by the fused enzyme, which then covalently modify proximal biomolecules. For instance, in enzyme-based systems, biotin ligases activate substrates to form reactive species like biotinoyl-5'-AMP that transfer the label to nearby lysine residues on proteins, while peroxidases generate radicals (e.g., biotin-phenoxyl radicals) that abstract hydrogens from amino acid side chains within a brief diffusion window. These labels allow for selective enrichment of tagged molecules, typically via affinity purification with streptavidin, followed by downstream analysis such as mass spectrometry to identify interactors. This principle exploits the nanoscale diffusion of reactive species to achieve specificity, distinguishing PL from global labeling methods and enabling the capture of dynamic, context-dependent associations in vivo. Non-enzymatic approaches, such as photoactivatable probes, operate on similar proximity-dependent tagging but rely on chemical catalysts rather than genetic fusions.^[2]^[7]^[6] The general workflow of proximity labeling begins with the genetic construction and expression of a fusion protein comprising the bait and labeling enzyme in the cellular system of interest. Upon addition of an exogenous substrate, the enzyme activates and initiates labeling, which occurs over minutes to hours depending on the system, after which cells are lysed to release labeled complexes. Tagged biomolecules are then purified using affinity handles like biotin-streptavidin interactions, and the enriched samples are subjected to proteomic or transcriptomic analysis for identification and quantification. This streamlined process supports high-throughput studies while minimizing artifacts from overexpression or fixation.^[2]^[7] A key feature of proximity labeling is its spatial resolution, governed by the diffusion distance and lifetime of the reactive intermediate, which confines tagging to a radius of approximately 5-20 nm—often around 10 nm in practice—comparable to the size of protein complexes. This nanoscale precision allows PL to delineate functional molecular neighborhoods, such as those in organelles or signaling hubs, and to detect transient interactions that evade conventional pull-down assays, thereby providing a robust tool for dissecting cellular architecture.^[2]^[6]^[7]

Comparison to Traditional Interaction Mapping

Traditional methods for mapping protein-protein interactions, such as co-immunoprecipitation (co-IP), primarily capture stable and high-affinity complexes but often fail to detect weak or transient interactions due to the requirement for cell lysis and purification, which can disrupt native cellular contexts.^[7]^[8] Yeast two-hybrid (Y2H) systems enable detection of binary interactions in vivo but are limited to proteins that can localize to the nucleus, lack native posttranslational modifications, and suffer from high rates of false positives, making them unsuitable for studying multiprotein complexes or interactions in their physiological environments.^[7]^[9] Techniques like Förster resonance energy transfer (FRET) and fluorescence lifetime imaging microscopy (FLIM) provide real-time measurement of protein proximities within a short distance range (typically <10 nm) but require fluorescent tagging and do not identify the specific protein identities involved, limiting their throughput and applicability to complex interactomes.^[7]^[8] Proximity labeling (PL) addresses these shortcomings by enabling the capture of weak and transient interactions directly in living cells, preserving native subcellular compartments and avoiding artifacts from overexpression or purification steps that plague traditional approaches.^[7]^[9] Unlike co-IP and Y2H, which depend on stable associations or artificial cellular relocations, PL techniques such as those using biotin ligase or peroxidase enzymes label proximal proteins in their endogenous context, facilitating the study of dynamic interactomes in real time without cell disruption.^[8] This in situ labeling extends to challenging targets like membrane-associated or insoluble proteins, which are often inaccessible to methods requiring solubility or nuclear trafficking.^[8] Despite these strengths, PL methods introduce potential drawbacks, including off-target labeling that can generate background noise and the necessity for genetic engineering to fuse labeling enzymes to bait proteins, which may alter protein folding or localization.^[7] These issues, while mitigable through optimized enzyme variants, contrast with the antibody-based simplicity of co-IP or the genetic screening ease of Y2H, though PL's native-context fidelity often outweighs such trade-offs for comprehensive mapping.^[9] Quantitatively, PL supports proteome-wide interaction profiling, routinely identifying hundreds of proximal proteins per bait—such as over 200 interactors in studies using APEX-based labeling—compared to the dozens typically recovered by co-IP, enabling broader discovery of novel associations and functional networks.^[7] This scale underscores PL's utility for high-throughput applications, surpassing the limited output of Y2H or the distance-constrained specificity of FRET in unraveling complex cellular interactomes.^[7]^[8]

Historical Development

Origins and Early Techniques

The concept of proximity labeling traces its roots to earlier techniques that utilized enzymes for localized modification of biomolecules, particularly in the context of electron microscopy and cell surface analysis. In the 1970s, horseradish peroxidase (HRP) emerged as a key tool for labeling cell surfaces and tracing neuronal pathways, where it was injected into tissues and detected via electron-dense reaction products to visualize uptake and transport in fixed samples.^[10] By the 1980s, HRP conjugation to antibodies enabled targeted peroxidase activity on live cell surfaces, facilitating the identification of extracellular matrix components and membrane proteins through enzymatic amplification.^[11] Complementing these enzymatic approaches, early biotinylation studies in the 1990s employed the biotin-avidin system to selectively label and isolate neural cell-surface proteins, allowing for their electrophoretic characterization and tracking in crude extracts without genetic engineering.^[12] These precursors laid the groundwork for proximity labeling by demonstrating the feasibility of enzyme-mediated or affinity-based modifications in situ, but they were limited to fixed cells, surface-exposed targets, or post-lysis analysis, lacking the ability to capture dynamic protein neighborhoods in living cells. The drive for dedicated proximity labeling arose from the need to map protein interactomes and organelle proteomes in vivo, overcoming the shortcomings of static methods like co-immunoprecipitation or yeast two-hybrid screens, which often missed transient or weak interactions in mammalian systems. Initial applications focused on eukaryotic cells to profile subcellular compartments, enabling the identification of proximal proteins under physiological conditions. A pivotal advancement came in 2012 with the development of APEX, an engineered ascorbate peroxidase derived from the dimeric enzyme in pea (Pisum sativum) or soybean (Glycine max), optimized for rapid, genetically encodable labeling in diverse cellular environments.^[13] APEX functions by oxidizing biotin-phenol substrates in the presence of hydrogen peroxide to generate reactive radicals that biotinylate nearby tyrosines within a 20 nm radius, allowing efficient labeling of proteins in mitochondria and the endoplasmic reticulum of live mammalian cells in under 1 minute. This method addressed prior limitations of HRP, such as its reliance on exogenous addition and poor activity in reducing cellular environments, providing a versatile tool for electron microscopy and proteomic profiling of organelle-specific proteomes.^[14] Concurrently in 2012, BioID was introduced as a complementary technique using a promiscuous mutant of the Escherichia coli biotin ligase BirA (BirA*), which promiscuously attaches biotin to nearby lysines within a ~10 nm radius without requiring cofactors beyond ambient biotin.^[15] Fused to nuclear proteins like lamin A, BioID enabled slower, steady-state labeling over 16-24 hours in living cells, capturing proximal interactors in the nuclear envelope and facilitating streptavidin-based purification for mass spectrometry analysis. Unlike APEX's rapid kinetics suited for acute labeling, BioID's ambient operation made it ideal for mapping stable protein associations in hard-to-access compartments, marking the first dedicated in vivo tools for unbiased interactome discovery in mammalian systems.^[16]

Evolution of Enzyme-Based Methods

Following the introduction of the original BioID method in 2012, which relied on a promiscuous biotin ligase requiring 18-24 hours for effective labeling but suffered from slow kinetics and high biotin concentrations, subsequent refinements focused on engineering smaller, more efficient enzymes to enhance speed, reduce off-target effects, and improve compatibility in diverse cellular environments.^[17] Parallel to these biotin ligase advancements, the peroxidase-based APEX system was evolved in 2015 into APEX2 through directed evolution using yeast surface display. This improved variant exhibits higher catalytic activity (up to 3-fold over APEX), better folding stability, and reduced expression requirements in mammalian cells, enabling more sensitive and robust proximity labeling with minimized background while maintaining the ~20 nm radius and rapid 1-minute kinetics.^[18] A key advancement came with BioID2 in 2016, an engineered variant of the Aquifex aeolicus biotin ligase reduced to approximately 27 kDa through deletion of non-essential domains, allowing better folding and localization in crowded cellular compartments like the nucleus.^[19] This smaller size enabled more selective fusion to bait proteins, required only 0.1 μM exogenous biotin for saturation (versus 1 μM for original BioID), and achieved robust proximity labeling within about 18 hours, thereby minimizing supplementation toxicity while maintaining a labeling radius of around 10 nm.^[19] These improvements made BioID2 particularly useful for studying nuclear and membrane-associated interactomes, as demonstrated in applications targeting nuclear envelope proteins where it outperformed the parent enzyme in specificity and efficiency.^[19] Further progress was driven by directed evolution techniques, culminating in TurboID and miniTurbo in 2018, both derived from the E. coli BirA biotin ligase through yeast display-based mutagenesis and selection for hyperactive variants.^[20] TurboID enabled rapid biotinylation in just 10 minutes using non-toxic biotin concentrations (50 μM), with a effective labeling radius of 5-10 nm, allowing capture of transient interactions that slower methods missed.^[20] Complementing this, miniTurbo—a 28 kDa truncated version lacking the N-terminal domain—offered similar 10-minute kinetics but with enhanced folding stability in dense cellular spaces, such as organelles, making it ideal for in vivo applications in model organisms like Drosophila and C. elegans.^[20] These enzymes dramatically expanded proximity labeling's temporal resolution, facilitating real-time mapping of dynamic protein complexes.^[20] To diversify beyond biotinylation and address background noise in mammalian systems, the PUP-IT system was developed in 2018, employing the prokaryotic enzyme PafA from Corynebacterium glutamicum to mediate pupylation—a covalent attachment of the small Pup tag (about 7 kDa) to proximate lysines on target proteins.^[21] Unlike diffusible biotin, Pup's immobility ensured low non-specific labeling, with effective tagging occurring over 6-24 hours under standard conditions, particularly suited for membrane protein interactomes like those of CD28 where it identified both known and novel partners with minimal background.^[21] This orthogonal approach provided an alternative to ligase-based methods, enabling parallel labeling strategies in complex proteomes.^[21] Building on these, split-enzyme variants emerged from 2017 onward to enable conditional activation, restricting labeling to induced protein associations and further refining specificity. Split-BioID, introduced in 2017, divides the BirA* ligase into two inactive fragments (split at residues E256/G257) that reassemble upon bait-prey dimerization, allowing validation of binary interactions while labeling vicinal proteins in 6-24 hours.^[22] Similarly, Split-TurboID (2020) adapts the hyperactive TurboID by fragmenting it into non-functional halves that complement upon contact, reducing overall labeling time to as little as 4 hours and enabling spatially precise mapping of organelle contact sites, such as endoplasmic reticulum-mitochondria interfaces, with reduced off-target effects compared to full-length enzymes.^[23] These conditional systems have proven invaluable for dissecting spatiotemporally regulated complexes, enhancing the method's utility in live-cell proteomics.^[23]

Enzyme-Based Methods

Biotin Ligase Techniques

Biotin ligase techniques in proximity labeling rely on engineered variants of the Escherichia coli biotin ligase BirA, particularly the promiscuous mutant BirA* (R118G), which is fused to a protein of interest to enable biotinylation of nearby proteins. The core mechanism involves the ligase catalyzing the formation of a reactive biotinoyl-5'-AMP intermediate from biotin and ATP, which diffuses briefly to covalently label proximal lysine residues on target proteins. This reaction proceeds as follows:

\text{Biotin} + \text{ATP} \xrightarrow{\text{BirA*}} \text{biotinoyl-5'-AMP} + \text{PP}_\text{i}

The short-lived intermediate limits labeling to proteins within approximately 10 nm of the fused ligase, providing spatial resolution for interaction mapping. The original BioID method utilizes a 35 kDa BirA* fusion that requires 18-24 hours of labeling in the presence of 50 μM exogenous biotin to achieve sufficient signal, though it suffers from high background due to self-biotinylation of the fusion protein itself. This extended labeling time captures stable interactions but can introduce artifacts from cellular changes over time, and the method's reliance on high biotin concentrations limits its use in some tissues. BioID's labeling radius is experimentally validated at approximately 10 nm, making it suitable for identifying proximal protein neighborhoods in mammalian cells.^[24] To address BioID's limitations, TurboID and miniTurbo were developed through directed evolution, incorporating the R118S mutation plus additional amino acid changes (e.g., E11K, N100Y, A124T) that enhance catalytic efficiency and reduce biotin dependence. These variants enable rapid labeling within 10 minutes at lower biotin concentrations (as low as 5 μM), minimizing toxicity and allowing capture of transient interactions without prolonged incubation. MiniTurbo, a truncated 28 kDa version with an N-terminal deletion, is particularly amenable to endogenous protein tagging via CRISPR, as its smaller size reduces steric hindrance in fusion constructs. Both exhibit a labeling radius similar to BioID but with 100-fold higher activity, enabling applications in diverse cellular contexts.^[25] Optimization of biotin ligase techniques has extended their utility to non-mammalian systems, including yeast and plants, where TurboID fusions successfully label proximal proteins under ambient temperatures and with adjusted biotin supplementation (e.g., 50 μM for 10-15 minutes in Arabidopsis protoplasts). To mitigate non-specific labeling from endogenous biotinylation or media contaminants, experimental controls typically include untagged cell lines or ligase-only fusions processed in parallel, ensuring that enriched proteins are proximity-dependent rather than artifactual. These adaptations have made the methods robust across eukaryotes while preserving the technique's hallmark of in vivo, covalent labeling.^[26]^[27]

Peroxidase Techniques

Peroxidase-based proximity labeling employs engineered ascorbate peroxidases, such as APEX, to catalyze the oxidative biotinylation of proximal biomolecules in living cells. The core mechanism involves the peroxidase oxidizing biotin-phenol in the presence of hydrogen peroxide (H₂O₂) to generate short-lived biotin-phenoxyl radicals, which covalently modify electron-rich residues like tyrosine and histidine on nearby proteins within approximately 20 nm. This process can be represented by the catalyzed reaction:

\text{Biotin-phenol} + \text{H}_2\text{O}_2 \rightarrow \text{biotin-phenoxyl radical} + \text{H}_2\text{O}

The original APEX enzyme, a 28 kDa monomeric variant derived from pea ascorbate peroxidase, was developed in 2012 primarily for electron microscopy but adapted for proteomic applications by 2013, enabling 1-2 minute labeling pulses. However, the requirement for H₂O₂ introduces cytotoxicity, limiting its use to short-term or fixed-sample labeling to minimize oxidative damage to cells.^[13]^[28] An improved variant, APEX2, incorporates the A134P mutation through directed evolution, enhancing catalytic activity by over fourfold compared to APEX while significantly reducing nonspecific background labeling and improving tolerance to H₂O₂ concentrations up to 2.5 mM. This allows for more efficient enrichment of proximal proteomes at lower enzyme expression levels, with reduced off-target noise in mass spectrometry analyses. Standard protocols for APEX2 typically involve preincubation with 1 mM biotin-phenol followed by brief pulses of 1 mM H₂O₂ for 1 minute to achieve optimal labeling while mitigating toxicity.^[18] These peroxidase techniques excel in capturing dynamic protein neighborhoods due to their sub-minute temporal resolution, making them suitable for studying transient interactions in living systems. Additionally, their compatibility with electron microscopy enables correlative light-electron imaging of labeled structures, providing spatial validation of proteomic data. Despite H₂O₂-related challenges, APEX and APEX2 have become widely adopted for mapping subcellular proteomes, such as mitochondrial matrices and endoplasmic reticulum membranes.^[28]^[18]

Alternative Enzyme Systems

PUP-IT (pupylation-based interaction tagging) represents an alternative enzyme-based proximity labeling approach derived from the prokaryotic pupylation pathway. In this system, the 54 kDa Pup ligase PafA is genetically fused to a bait protein, enabling it to covalently attach a modified Pup(E) tag—a 64-amino-acid peptide—to lysine residues on proximal prey proteins within a labeling radius of less than 10 nm.^[29] The process requires 6-24 hours for efficient tagging and is fully genetically encoded, eliminating the need for exogenous substrates.^[29] Unlike diffusion-prone methods such as BioID, PUP-IT's direct ligation of the larger Pup tag minimizes background noise by preventing tag dissemination beyond the immediate vicinity.^[29] Split enzyme systems extend proximity labeling by incorporating conditional activation, where enzyme reconstitution depends on bait-prey interactions to initiate tagging. Split-BioID divides the BirA* biotin ligase into two inactive fragments, each fused to a separate bait; dimerization or complex formation reassembles the enzyme, triggering biotinylation of nearby proteins and enabling validation of transient interactions.^[22] Building on faster variants like TurboID, Split-TurboID similarly splits the promiscuous TurboID enzyme, allowing contact-dependent labeling within 4-16 hours to map dynamic protein complexes with reduced off-target effects.^[23] Less common alternatives leverage engineered enzymes for diverse tagging modalities. For instance, the NEDDylator fuses an E2-conjugating enzyme (Ubc12) to a bait, catalyzing proximity-dependent neddylation—a ubiquitin-like modification—on lysine residues of nearby proteins, providing an orthogonal tag for interactome analysis.^[30] Engineered dehalogenases, such as modified haloalkane dehalogenases, enable covalent attachment of alternative synthetic tags to proximal sites but are primarily used for targeted labeling rather than comprehensive mapping. These systems expand tag variety while maintaining enzymatic specificity, though their adoption lags behind more established biotin- or pupylation-based tools. More recent ubiquitin-based systems, such as those for selective E3 ligase substrate tagging, further diversify proximity labeling options.^[31]

Non-Enzymatic Methods

Chemical Proximity Labeling

Chemical proximity labeling encompasses non-enzymatic strategies that utilize small-molecule probes to selectively tag biomolecules in close spatial proximity to a target, typically through abiotic reactive groups activated by UV light or chemical means. These probes are directed to specific sites via conjugation to antibodies, ligands, or other targeting moieties, enabling substrate-driven labeling without the need for genetic fusion of enzymes to the protein of interest. A prominent class involves diazirine-based photo-crosslinkers, which are incorporated into biotinylated probes for subsequent enrichment and identification of labeled interactors. This approach is particularly suited for mapping interactions in native cellular environments, as it avoids perturbations associated with overexpression systems.^[32] The core mechanism relies on the photoactivation of diazirine moieties, where UV irradiation (typically at 365 nm) cleaves the three-membered ring, expelling nitrogen gas and generating a singlet carbene intermediate. This highly electrophilic species rapidly forms covalent bonds by inserting into nearby C-H, N-H, S-H, or O-H bonds on amino acid side chains or backbones, with a labeling radius of approximately 1-5 nm determined by the probe's linker length and the carbene's short lifetime (on the order of nanoseconds). While this provides nanoscale resolution for capturing transient or weak interactions, the carbene's indiscriminate reactivity can lead to higher background labeling compared to more selective enzymatic methods, often mitigated by optimizing exposure times and using quenchers like water or amines.^[33] Alternative chemical activation, such as through aryl azides generating nitrenes, follows a similar principle but may involve triplet states for broader reactivity.^[34] Representative examples include antibody-conjugated diazirine-biotin probes for labeling surface protein interactomes, where the antibody binds a target receptor on live cells, and UV activation cross-links adjacent membrane proteins for streptavidin pulldown and mass spectrometry analysis. This has been applied to profile neighborhoods around receptors like EGFR without cellular engineering.^[35] Another approach leverages chemical inducers of proximity (CIP) systems, such as CATCHFIRE (chemically assisted tethering of chimera by fluorogenic-induced recognition), developed in 2023, which uses small-molecule "match" inducers to reversibly dimerize genetically tagged proteins (via a 11-residue peptide and a 114-residue domain). This system's reversibility, achieved by inducer washout, allows temporal control and imaging of dynamic interactions in living cells.^[36] These methods offer key advantages, including compatibility with primary cells, tissues, and non-transfected systems, bypassing the limitations of genetic manipulation while preserving endogenous expression levels and stoichiometry. However, challenges like non-specific labeling necessitate rigorous controls, such as no-probe or no-UV conditions, to ensure specificity. Overall, chemical proximity labeling complements affinity purification by capturing proximity-based associations in situ, providing insights into undiscovered interactomes.^[37]

Photocatalytic Proximity Labeling

Photocatalytic proximity labeling utilizes light-activated catalysts to generate reactive species that covalently tag biomolecules within a localized radius, typically 10-100 nm, enabling non-genetic spatial proteomics without enzymatic dependencies. Common photocatalysts include transition metal complexes such as ruthenium (Ru) and iridium (Ir) complexes, alongside organic dyes like thiorhodamine, which upon visible light irradiation—often deep-red at 660 nm for enhanced tissue penetration—produce reactive oxygen species (ROS) like singlet oxygen or triplet intermediates. These reactive species facilitate bioconjugation with tags such as azide-biotin, allowing downstream enrichment and analysis via click chemistry.^[38] The core mechanism proceeds from light excitation of the photocatalyst, yielding ROS or short-lived radicals that diffuse briefly to label proximal proteins before deactivation, providing spatiotemporal precision. In the seminal μMap-Red method (2022), a Sn(IV) chlorin e6 photocatalyst activated by 660 nm light generates triplet nitrenes from phenyl azide biotin probes, achieving ~10 nm resolution for mapping protein environments in cancer cells and tissues with low phototoxicity. This non-genetic approach minimizes cellular perturbation compared to enzyme-based systems and suits non-transfectable samples like primary cells.^[39]^[38] Notable methods include CAT-S (2024), a bioorthogonal system employing an Ir photocatalyst and thio-quinone methide probe under blue light to label mitochondrial proteomes in primary cells and tissues, such as human T cells and mouse kidney, with 61-87% specificity and no genetic engineering required. PNPL (2025), a nanoparticle-based innovation, leverages photocatalytic nanoscale metal-organic frameworks (pnMOFs) to produce singlet oxygen for histidine-selective biotinylation, targeting subcellular locales like the ER in living HeLa cells.^[40]^[41] Advancements in temporal control feature time-resolved variants like CAT-ER (2025), which uses an ER-targeted Ir photocatalyst and thio-quinone methide for in situ profiling of proteome dynamics during endoplasmic reticulum stress, identifying regulators such as NFIP2 in UPR-to-apoptosis shifts across cell types including Jurkat and RAW264.7. BRET-activated platforms, exemplified by BRET-ID (2025), couple NanoLuc luciferase to a photosensitizer for bioluminescence-triggered ROS generation, enabling in vivo spatial mapping of interactomes—like G3BP1 in mouse tumor xenografts—without external illumination and with 1-minute resolution. These developments underscore photocatalytic labeling's edge in vivo compatibility and reduced invasiveness over traditional enzyme methods.^[42]

Applications

Protein Interactome Analysis

Proximity labeling (PL) has emerged as a powerful tool for mapping protein interactomes by capturing both stable and transient protein-protein interactions in living cells, overcoming limitations of traditional affinity purification methods that often miss weak or dynamic associations. By fusing labeling enzymes to bait proteins, PL biotinylates proximal preys, enabling their enrichment and identification via mass spectrometry (MS). This approach is particularly valuable for complex macromolecular assemblies, where it reveals neighborhood relationships without disrupting native cellular environments. In nuclear pore complex (NPC) studies, BioID and TurboID have identified extensive interactomes, with BioID applied to multiple nucleoporins revealing 47–94% of labeled candidates as NPC-related proteins across baits like Nup160 and Nup53, encompassing over 200 proximal proteins in aggregate datasets. Similarly, APEX has been used to map chromatin-associated proteomes, such as in dCas9-APEX fusions targeting promoters like hTERT and c-MYC, identifying dozens of proximal factors involved in transcription and DNA maintenance. These use cases demonstrate PL's ability to delineate multi-protein networks in structured cellular compartments.^[43] The typical workflow integrates PL with MS for bait-prey identification: cells expressing enzyme-bait fusions are labeled with biotin substrates, followed by lysis, streptavidin pulldown, on-bead digestion, and LC-MS/MS analysis to quantify biotinylated peptides. High-confidence hits are filtered using abundance ratios (e.g., bait/control >2-fold) and statistical models like SAINT, distinguishing specific interactors from background. This process yields binary interaction networks, often visualized as graphs to highlight core modules. Notable examples include the EGFR interactome, where AirID-based PL in cancer cells identified 22 extracellular proximal proteins, including 12 known partners like integrins, revealing ligand-dependent dynamics in signaling pathways. In plants, BioID screening near the transcription factor OsFD2 in rice protoplasts identified 62 high-confidence neighbors, validated by BiFC and Y2H for key candidates like Q5VMJ3 (profilin LP04). Quantitative metrics underscore reliability: false discovery rates below 5% are achieved with biological replicates and SAINT scoring (≥0.8 threshold), while PL captures weak affinities below 1 μM by labeling based on spatiotemporal proximity rather than binding strength. The ~10 nm labeling radius ensures specificity for true interactors.

Subcellular and Organelle Mapping

Proximity labeling (PL) techniques enable the profiling of proteomes within specific subcellular compartments by fusing labeling enzymes to organelle-targeting signals, allowing for the biotinylation and subsequent identification of proximal proteins. This approach has been particularly effective with APEX variants, which facilitate rapid labeling in membrane-enclosed spaces such as the endoplasmic reticulum (ER) lumen and mitochondrial matrix. For instance, APEX-mediated labeling in the ER has identified promoters of ER-mitochondrial contacts by coupling proximity biotinylation with biochemical fractionation, revealing key interactors at these interfaces. Similarly, APEX2 has been used to map the proteome of the cytosol-facing surfaces of outer mitochondrial and ER membranes, enriching for compartment-specific proteins without requiring cell lysis. In peroxisomes, APEX-based methods have profiled peroxisomal proteomes, identifying established and novel constituents, highlighting the technique's utility in organelle-specific discovery. Recent innovations like iAPEX have improved resolution by reducing background labeling in such mappings.^[44] TurboID, a high-efficiency biotin ligase, complements these applications by capturing dynamic processes at the plasma membrane. Fusions of TurboID to membrane-associated proteins, such as eisosome components in yeast, have identified proximal interactors involved in membrane organization and dynamics, enabling the study of transient associations under physiological conditions. This method's speed—labeling within minutes—makes it suitable for tracking plasma membrane proteome changes in response to stimuli, such as in neuronal signaling. To achieve precise organelle targeting, PL enzymes are genetically fused to signal peptides or domain-specific anchors that direct localization to desired compartments. For example, mitochondrial targeting sequences guide APEX to the matrix, while ER signal peptides direct labeling to luminal proteomes, ensuring spatial restriction of the biotinylation radius to 10-20 nm. Validation of these localizations often involves correlation with electron microscopy (EM), where APEX-generated electron-dense deposits confirm enzyme positioning at high resolution, as demonstrated in yeast organelle markers. This integration enhances confidence in compartment-specific enrichments by aligning proteomic data with ultrastructural observations. Notable examples illustrate PL's impact in complex cellular contexts. Recent studies using proximity labeling have mapped synaptic proteomes in murine brain neurons, identifying hundreds to thousands of compartment-specific proteins across presynaptic, postsynaptic, and glial interfaces, revealing specialized functional signatures. For non-membrane-bound structures, split-APEX variants have enabled RNA-granule mapping by reconstituting the enzyme only upon granule assembly, labeling proximal proteins and RNAs in stress granules and P-bodies to uncover regulatory networks. The resolution of PL in subcellular mapping stems from its ability to enrich compartment-specific proteomes while distinguishing soluble from membrane-associated fractions. APEX2 and TurboID fusions to cytosolic or nuclear signals yield distinct profiles, with membrane-targeted variants enriching integral membrane proteins over cytosolic ones by factors of 5-10-fold, as shown in comparative labeling of HEK293 cells. This selectivity allows differentiation between cytosol and membrane proteomes, providing insights into compartmental organization without extensive fractionation.

Biomedical and Organism-Specific Uses

Proximity labeling has been instrumental in elucidating protein interactomes in cancer, particularly for oncogenic mutants like KRAS. In a 2025 study, TurboID-based proximity labeling integrated with quantitative proteomics mapped the interactomes of wild-type KRAS and mutants G12C, G12D, and G12V, identifying over 1,000 proximal proteins and revealing mutant-specific interactions that inform targeted therapies for KRAS-driven cancers.^[45] In neuroscience, a 2025 review highlighted applications of BioID, APEX, and HRP in profiling synaptic proteomes under pathological conditions, including Alzheimer's disease, and their potential for identifying dysregulated proteins and biomarkers in amyloid-beta-induced neurodegeneration.^[46] Beyond mammalian systems, proximity labeling facilitates organism-specific studies in plants, microbes, and model organisms. In Arabidopsis thaliana, pupylation-based proximity labeling (PUP-IT) in 2025 identified regulatory factors in cellulose biosynthesis, such as BEN1 that controls cellulose synthase complex trafficking via ubiquitination during cell wall formation.^[47] For microbial interactions, proximity labeling mapped SARS-CoV-2-host factors in 2021, identifying 437 human proteins proximal to viral proteins via BioID and split-BioID, providing insights into viral replication mechanisms.^[48] In zebrafish, TurboID-enabled proximity labeling in 2021 profiled local proteomes during embryonic development, revealing dynamic protein networks essential for organogenesis and regeneration.^[49] Proximity labeling also aids viral and pathogen research by pinpointing host factors near viral proteins to discover antiviral targets. For instance, super-resolution proximity labeling in 2023 delineated anti-viral protein networks interacting with SARS-CoV-2 ORF3a and M proteins, validating RNF5 as a host E3 ligase that reduces viral infectivity.^[50] Emerging applications include hybridization-proximity labeling (HyPro) for mapping RNA-protein networks in primary tissues. A 2025 enhancement of HyPro technology enabled in situ profiling of protein interactomes around endogenous RNAs in live cells and tissues, identifying spatially resolved networks that bridge transcriptomics and proteomics in complex biological contexts.^[51]

Challenges and Advances

Technical Limitations

Proximity labeling techniques, while powerful for mapping protein interactions, face several implementation challenges that can introduce artifacts and confound results. In biotin ligase-based methods like BioID, background labeling arises from the enzyme's self-biotinylation and the presence of endogenously biotinylated proteins, such as mitochondrial carboxylases, which can appear as false interactors in downstream analyses.^[52] Similarly, peroxidase-based approaches like APEX require hydrogen peroxide to activate labeling, which induces oxidative stress and cytotoxicity, particularly limiting their use in vivo or in sensitive cell types.^[52] Genetic fusion of the labeling enzyme to the bait protein can also disrupt native localization, folding, or function, potentially altering the captured interactome.^[7] Analytical interpretation of proximity labeling data, often coupled with mass spectrometry, is hampered by high rates of false positives due to nonspecific binding during streptavidin enrichment and incomplete separation of labeled from unlabeled peptides.^[52] Labeling exhibits inherent biases, preferentially targeting surface-exposed lysine residues in BioID/TurboID systems or tyrosines in APEX, which may underrepresent proteins lacking these amino acids and skew proteome coverage.^[52]^[7] The spatial resolution of proximity labeling is constrained to a 5–20 nm radius due to the diffusion of reactive intermediates, which captures proteins in close proximity but not necessarily in direct physical contact, thus blurring fine-scale mapping of molecular interfaces.^[52] Temporal resolution varies significantly across methods; for instance, BioID requires 12–24 hours for effective labeling, suitable only for stable interactions, whereas TurboID achieves robust labeling in as little as 10 minutes, enabling capture of more dynamic associations.^[2]^[53] To address these limitations, researchers employ controls such as label-free bait proteins or enzyme-only fusions to subtract background signal, alongside computational tools like SAINT scoring, which uses quantitative mass spectrometry data from negative controls to assign confidence scores and filter false positives with false discovery rates below 1%.^[52]^[54]

Recent Innovations and Future Directions

Recent innovations in proximity labeling (PL) have focused on enhancing specificity, spatiotemporal control, and applicability to complex biological systems. In 2024, the POST-IT system was introduced as a non-diffusive proximity tagging method utilizing pupylation via PafA and HaloTag ligands to identify drug targets in live cells and organisms, such as revealing SEPHS2 as a dasatinib target in HEK293T cells and zebrafish embryos.^[55] This approach minimizes off-target labeling, enabling precise drug screening by capturing targets in native contexts without genetic modification of the bait.^[55] Advancements in photocatalytic PL have improved temporal resolution for dynamic processes. A 2025 development, CAT-ER, employs an ER-targeted iridium photocatalyst with a thio-quinone methide probe to profile endoplasmic reticulum (ER) proteome changes during unfolded protein response (UPR) to apoptosis transition in cancer cells, identifying regulators like NFIp2 and EMC2 without genetic manipulation.^[42] This time-resolved method achieves high ER specificity across diverse cell types, including HeLa and Jurkat cells, by light-triggered labeling.^[42] Enzyme engineering has boosted sensitivity in RNA-associated PL. The 2025 enhanced HyPro2 variant, with mutations D14K and K112E, optimizes biotinylation conditions (50% trehalose, 5 μM hemin) to map interactomes of single RNA molecules, such as C9orf72 transcripts, outperforming prior methods in detecting low-abundance compartments with fewer than 10 RNAs.^[51] This re-engineering reduces diffusion and enhances peroxidase activity, facilitating proteomics of compact RNA structures like ACTB transcription sites.^[51] New variants address in vivo challenges and reversibility. Photocatalytic nanoscale metal-organic frameworks (pnMOFs) enable PNPL in 2025 by generating singlet oxygen to label histidine residues near nanoparticles, mapping ~7,200 sites in live HeLa cells and supporting applications in tumor-targeted photodynamic therapy.^[56] Light-controlled photocatalytic systems, such as those in CAT-ER, offer reversibility through on-off illumination, allowing termination of labeling to study transient interactions.^[42] Looking ahead, PL is poised for integration with single-cell mass spectrometry to resolve heterogeneous interactomes in tissues, as suggested by recent proteomics pipelines emphasizing subcellular specificity.^[57] Artificial intelligence models are emerging to predict interactomes from PL data, enhancing interpretation of large-scale PPI networks by distinguishing positive and negative interactions via latent space representations.^[58] Expansion to non-protein biomolecules includes adaptations for lipids via singlet-oxygen-based POCA and glycans through photoaffinity strategies, broadening PL to glycan-protein interactions.^[59] Overall trends favor non-genetic, light-inducible methods like photocatalysis for clinical translation, enabling spatiotemporal control in vivo without genomic alterations.^[60]

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