TurboID Proximity Labeling Services for Interactome Discovery

TurboID has ~100× higher catalytic activity than BioID (kcat ~0.3 s⁻¹ vs. ~0.003 s⁻¹), reducing required labeling time from 18–24 hours to 10–60 minutes. This matters for three reasons: (1) transient and weak interactions — which may exist for only minutes — are captured before they dissociate, (2) low-abundance or toxic bait proteins can be used because the cells only need to survive a 10-minute biotin pulse rather than overnight overexpression, and (3) temporal resolution — the interactome can be sampled at multiple time points after a stimulus (e.g., drug treatment, signaling activation), which is impossible with the 18-hour averaging window of BioID. Side-by-side comparisons by Branon et al. (2018) showed TurboID produced stronger biotinylation signal in 10 minutes than BioID did in 18 hours.

Choose miniTurbo (28 kDa) over full-length TurboID (35 kDa) when: (1) your bait protein is small or compact — a 35 kDa fusion tag may sterically hinder folding, localization, or interactions, (2) your bait functions in a confined subcellular compartment (e.g., mitochondrial intermembrane space, peroxisome lumen) where tag size is critical, or (3) you observe mislocalization or aggregation of the TurboID fusion construct during expression validation. miniTurbo has comparable labeling kinetics to TurboID with marginally lower background in some compartments. If tag size is not a concern and maximum labeling efficiency is the priority, full-length TurboID is preferred.

Full-length TurboID labels all proteins within ~10 nm of the bait — but cannot distinguish a protein that physically contacts the bait from one that merely resides in the same subcellular compartment (a "bystander"). Split-TurboID divides the enzyme into two inactive fragments (N-Turbo and C-Turbo), each fused to a different bait protein. Active enzyme reconstitutes only when the two bait proteins are within ~10 nm of each other — and only the proteome at that interface is biotinylated. Cho et al. (2022) applied this to ER–mitochondria contact sites, identifying 101–115 proteins specifically enriched at the interface that bulk organelle labeling missed. This makes Split-TurboID the method of choice when you need to confirm whether two proteins of interest physically interact, rather than simply reside in the same organelle.

Yes — and this is one of TurboID's most powerful capabilities. Unlike APEX2, which requires toxic H₂O₂ for labeling, TurboID uses endogenous biotin — a natural vitamin (B7) — as its substrate. For mouse models, biotin is administered via drinking water (0.5 mg/mL) or intraperitoneal injection for 7–14 days, and AAV vectors deliver TurboID fusions to specific tissues or cell types. Wei et al. (2023) mapped cell-type-specific secretomes across 21 tissues in live mice using this approach — identifying exercise-regulated secreted proteins that no cell-culture-based method could discover. Published in vivo applications include mouse (liver, brain, muscle), Arabidopsis, rice, C. elegans, Drosophila, and zebrafish.

A standard TurboID experiment requires 2 × 10⁷ cells per condition. Essential controls include: (1) TurboID-only (no bait fusion) — expressed at comparable levels to the bait fusion to subtract non-specific biotinylation, (2) no-biotin control — identifies endogenous biotinylated proteins and streptavidin-binding contaminants, and (3) biological triplicates for SAINT statistical scoring. For organelle-targeted TurboID, a localization-matched control (e.g., cytosolic TurboID for a mitochondrial-targeted experiment) is recommended. These controls are not optional — they are the difference between a high-confidence interactome and a list of non-specific hits. Our experimental design consultation ensures the appropriate control set is planned before the first experiment.

AP-MS enriches bait complexes from cell lysates under native (non-denaturing) conditions. This works well for stable, high-affinity complexes but systematically loses weak/transient interactions (which dissociate during lysis and washing) and membrane protein complexes (which require detergents that may disrupt native interactions). TurboID applies the biotin tag covalently before lysis — weak interactors are captured in situ, membrane proteins are solubilized by denaturing lysis without disrupting pre-existing interactions, and stringent denaturing washes during enrichment eliminate post-lysis background. The trade-off: AP-MS identifies direct complex members with high confidence if the complex is stable; TurboID captures the full proximal proteome but requires SAINT/CRAPome filtering to distinguish specific from non-specific proximity. In practice, the two methods are complementary — AP-MS for stable core complexes, TurboID for the broader interaction neighborhood including transient and membrane-proximal partners.