What Is Biotinylation? How Biotin Is Attached to Peptides

Published October 2026 · Patriot Labs Research Team · ~22 min read

The short answer

Biotinylation is the covalent attachment of biotin — a small vitamin, molecular weight 244.31 g/mol — to a peptide, protein or other molecule, so that the molecule can be captured or detected through biotin’s exceptionally tight binding to the proteins streptavidin and avidin. That binding is the strongest known non-covalent protein–ligand interaction, with a dissociation constant near 10⁻¹⁴ molar, which means a biotinylated molecule can be pulled out of a complex mixture, pinned to a surface, or lit up for detection with almost no leakage. Biotin is small enough that the tag rarely disturbs what it is attached to, and it can be added chemically or by an enzyme.^[1][2]

That one pairing — a tiny, easily attached tag and a protein that grips it like a vice — is why biotinylation turns up everywhere in the research literature. The rest of this guide takes it apart: what biotin is and why streptavidin holds it so tightly; the chemical reagents that bolt biotin onto amines, thiols or carboxyls; the spacer arms and water-soluble variants that make the tag reachable; the enzymatic route that places a single biotin at one defined site; the proximity-labeling methods (BioID and TurboID) that turned the enzyme into a tool for mapping protein neighbourhoods; what biotinylated peptides are actually used for; and how a biotinylated peptide is told apart from its parent in the analytical laboratory.

What biotinylation actually is

Biotinylation means covalently attaching biotin to another molecule. Biotin itself is vitamin B7 (historically also called vitamin H or coenzyme R), a small bicyclic compound of 244.31 g/mol made of a fused ureido ring and a tetrahydrothiophene ring, with a five-carbon valeric-acid side chain ending in a carboxyl group.^[1] That carboxyl is the handle: almost every biotinylation reagent is built by taking biotin and converting its valeric-acid tail into a reactive group, or extending it with a linker that carries one.

The reason to add such a tag is that biotin is recognised with extraordinary specificity and strength by a family of binding proteins — avidin from egg white, streptavidin from bacteria, and engineered relatives. Attaching biotin therefore gives an otherwise unremarkable peptide a universal docking point: anywhere a streptavidin surface, bead or reporter is waiting, the biotinylated molecule will bind and stay bound. Because biotin is small relative to a protein, the modification is “rapid, specific and is unlikely to disturb the natural function” of the labeled molecule, and more than one biotin can be attached to a single target so that several streptavidin molecules can dock at once, amplifying a signal.^[1]

There are two broad ways to attach it, and the distinction organises the whole field. Chemical biotinylation uses a reactive biotin reagent that couples to a chemical group already present on the target — usually amines, but also thiols, carboxyls or oxidised sugars — and because those groups appear many times on a typical protein, chemical labeling is fast but gives a spread of biotins at poorly controlled positions.^[1] Enzymatic biotinylation uses a bacterial enzyme, biotin ligase, to attach a single biotin to one specific lysine inside a short recognition sequence, giving a defined number at a defined site.^[1][5] Everything that follows is a variation on these two themes.

Biotin and the streptavidin bond: why it works

Biotinylation is only useful because of what biotin binds to, so the binding proteins come first. The biotin–streptavidin interaction is a non-covalent bond that, in the words of the literature, “approaches the strength of a covalent bond” and is far stronger than most antibody–antigen pairings.^[1] Streptavidin, a 52-kilodalton tetramer from the bacterium Streptomyces avidinii, binds biotin with a dissociation constant on the order of 10⁻¹⁴ molar; avidin, the egg-white protein, binds free biotin even harder, around 10⁻¹⁵ molar.^[2][3] To put that in perspective, the complex survives organic solvents, the denaturant guanidinium chloride, detergents such as SDS and Triton, proteases, and extremes of temperature and pH; breaking it usually requires something like 6–8 molar guanidinium chloride at pH 1.5, which also destroys most proteins.^[1][2]

The structural reason was settled in 1989, when two groups independently solved the crystal structure of streptavidin with biotin bound.^[9][10] Each streptavidin subunit folds into an eight-stranded antiparallel β-barrel, and biotin sits in a pocket at one end of the barrel. Three things combine to produce the enormous affinity: a dense network of roughly eight hydrogen bonds to the biotin’s ureido and carboxyl groups; a deep hydrophobic pocket lined with tryptophan residues, including one tryptophan contributed by a neighbouring subunit; and a flexible surface loop (often called L3/4) that folds down over the bound biotin like a lid, dramatically slowing its escape.^[2] The affinity is greater than the hydrogen bonds alone would predict, because the closed, rigid, water-excluding pocket makes dissociation extraordinarily slow.

The three binding proteins differ in ways that matter in practice. Avidin is a glycoprotein — about a tenth of its mass is carbohydrate — and it is strongly basic, which makes it bind many surfaces non-specifically and raise background. Streptavidin carries no sugar and has a near-neutral isoelectric point, so it binds much more cleanly. NeutrAvidin is avidin with its sugars removed, which lowers its charge and background while keeping high biotin affinity. A fourth complication is that streptavidin carries an exposed RYD sequence that mimics the RGD cell-adhesion motif, which can cause it to stick to integrin-bearing cells; this is one more reason assays are designed carefully.^[2] The table below summarises the differences.

Engineered variants extend the toolkit further. Monovalent streptavidin presents a single functional biotin site on an otherwise intact tetramer, so it binds a biotinylated target with full 10⁻¹⁴-molar affinity without cross-linking several of them together — useful for tracking single receptors.^[8] Traptavidin is a mutant with roughly ten-fold slower biotin dissociation and greater mechanical and thermal stability, for applications where the bond must never let go.^[11] These are reminders that “streptavidin” is really a platform of related proteins chosen to fit the experiment.

Chemical route I: amine-reactive NHS esters

The most common way to biotinylate a peptide or protein chemically is through its primary amines, and the dominant reagent is an NHS-ester biotin. Primary amines are abundant and reactive: they occur on the ε-amino group of every lysine side chain and once more at the free α-amino group of the N-terminus.^[1][16] An NHS ester (N-hydroxysuccinimide ester) is an activated form of biotin’s carboxyl that reacts with a deprotonated amine to form a stable amide bond, releasing the N-hydroxysuccinimide group as a leaving group.

Because it is the uncharged, deprotonated amine that attacks, the reaction is run at mildly basic pH, roughly 7 to 9, and typically at 4–37 °C for anywhere from a few minutes to overnight.^[16] Two practical points follow. First, the NHS ester competes with its own hydrolysis by water: the half-life of hydrolysis is only about two to four hours at pH 7 and falls to a few minutes at pH 9, so the reagent is dissolved immediately before use and the reaction is not left longer than needed.^[16] Second, amine-containing buffers such as Tris and glycine will consume the reagent and must be avoided; a phosphate or bicarbonate buffer is used instead.

The trade-off of amine labeling is control. A protein such as serum albumin has dozens of lysines — roughly 59 primary amines, of which 30–35 are on the surface — so NHS-biotin produces a population of molecules carrying biotin at varying positions and in varying numbers.^[16] For many detection assays that heterogeneity is acceptable or even helpful, because multiple biotins amplify the streptavidin signal. When a single, defined attachment is needed, chemists turn to the thiol route or to enzymatic labeling described below. For synthetic peptides specifically, the cleanest approach is to incorporate the biotin during solid-phase synthesis — coupling a biotin building block, or biocytin (a biotin joined to a lysine), to the N-terminus on the resin — so that every molecule carries exactly one biotin at one end. The chemistry of building peptides this way is covered in What Is Solid-Phase Peptide Synthesis?

Chemical route II: thiol, carboxyl and carbonyl

When amine labeling is too indiscriminate, other functional groups offer more selective handles.

Thiol-reactive (cysteine) biotin

The sulfhydryl (–SH) group of a free cysteine is far less common than lysine on most proteins, which makes it an excellent site-specific target. The standard reagent is a maleimide-biotin; maleimides react roughly a thousand times faster with a free thiol than with an amine at neutral pH, and the reaction is run at pH 6.5–7.5.^[16] Haloacetyl (iodoacetyl) and pyridyl-disulfide biotins are alternatives, the last of which forms a cleavable disulfide that can later be released with a reducing agent such as DTT. Because a synthetic peptide can be made with exactly one cysteine placed wherever the designer wants, maleimide chemistry is a favourite route to a single, oriented biotin on a peptide. Disulfide bonds must first be reduced to expose the thiol, and if a protein has no usable cysteine, a lysine can be converted to a thiol first with a reagent such as 2-iminothiolane (Traut’s reagent).^[1][16]

Carboxyl-reactive biotin

Biotin can also be attached to carboxyl groups — the C-terminus and the side chains of aspartate and glutamate — using an amine-functionalised biotin together with the carbodiimide crosslinker EDC. This reaction runs at acidic pH, about 4.5 to 5.5, in a buffer such as MES, and avoids amine- and carboxyl-containing buffers.^[16] It is less common than amine or thiol labeling because carboxyls are abundant and the chemistry is harder to control, but it is useful when amines must be preserved.

Carbonyl (glycan) and photoreactive biotin

Glycoproteins offer a route that targets their sugars rather than their polypeptide: mild periodate oxidation converts cis-diols on sialic acids into aldehydes, which then react with a hydrazide- or alkoxyamine-biotin to form a hydrazone. Because the sugars of an antibody sit away from its antigen-binding region, this is a popular way to biotinylate antibodies without blocking their function.^[1][16] Finally, photoreactive biotin carries an aryl-azide group that is chemically inert until ultraviolet light converts it into a highly reactive nitrene, which inserts into almost any nearby bond. This is used when a molecule has no convenient amine, thiol, carboxyl or sugar, or when labeling must be triggered at a chosen moment.^[1]

Spacer arms, sulfo-reagents and reversible biotin

Choosing the reactive chemistry is only half the decision; the linker between that reactive group and the biotin matters just as much. The biotin pocket of streptavidin sits roughly nine angstroms below the protein’s surface, so a biotin attached hard against a large molecule can be partly hidden, and streptavidin binds it poorly.^[16] The fix is a spacer arm: a short linker that holds the biotin out where it is accessible. Reagent catalogues are organised around arm length — a plain NHS-biotin has a spacer of about 13.5 Å, a “long-chain” (LC) version about 22.4 Å, an “extra-long” (LC-LC) version about 30.5 Å, and a PEG4 version about 29 Å — and the general rule is that “long spacer arms reduce steric hindrance and result in enhanced interaction of avidin and biotin.”^[16]

A second axis is water solubility. Plain NHS-esters are hydrophobic and must first be dissolved in an organic solvent such as DMSO or DMF before dilution into the aqueous reaction; being uncharged, they can also cross cell membranes and label the inside of cells. Sulfo-NHS esters carry a charged sulfonate group on the NHS ring that makes them water-soluble and membrane-impermeant, so they are the standard choice for labeling only the outside surface of intact cells.^[16] PEG spacers (polyethylene-glycol linkers) add both length and lasting water solubility, which is why reagents such as NHS-PEG4-biotin are popular for antibodies and for keeping a labeled protein soluble.

Finally, the very strength of the biotin–streptavidin bond is sometimes a problem: once a protein is captured, it can be hard to get back without destroying it. Several tricks make the capture reversible. A weaker biotin analogue, desthiobiotin, binds streptavidin firmly enough to capture but loosely enough to be eluted with a gentle excess of free biotin. A pH-switchable analogue, iminobiotin, binds avidin or streptavidin strongly at alkaline pH and releases near pH 4, allowing mild recovery.^[1] And for some purposes the bond itself can be broken reversibly: a short incubation in water above 70 °C has been shown to release biotinylated DNA from streptavidin without denaturing the protein, so the support can be reused.^[15]

Enzymatic biotinylation: BirA and the AviTag

Enzymatic biotinylation solves the control problem completely: it puts one biotin on one known residue and nothing else. The enzyme is BirA, the Escherichia coli biotin ligase (biotin holoenzyme synthetase), a roughly 35-kilodalton protein whose natural job is to attach biotin to a single lysine on the biotin carboxyl carrier protein of acetyl-CoA carboxylase, a step cells need to make fatty acids.^[1] BirA works in two steps at one active site: first it activates biotin with ATP to form a high-energy mixed anhydride, biotinyl-5′-AMP; then it transfers the biotinyl group onto the ε-amino group of the target lysine, forming a stable amide bond and releasing AMP.^[1]

The breakthrough that turned this into a research tool was discovering that BirA’s long natural substrate could be shrunk to a tiny peptide. Peptide-library screening identified a short consensus sequence that BirA will biotinylate, and this was refined into the 15-residue AviTag (also called the acceptor peptide, AP), with the sequence GLNDIFEAQKIEWHE; BirA attaches biotin to the single lysine in that sequence.^[1][5][6] A protein is genetically fused to the AviTag at its N-terminus, C-terminus or an accessible loop, then incubated with BirA, biotin and ATP — or co-expressed with BirA inside a cell — and the result is one biotin at one defined position, with no other protein modified.^[1] That site-specificity gives a uniform, oriented reagent, which is why AviTag biotinylation is used to build MHC tetramers and other precisely-assembled affinity reagents.

The approach has been extended in clever ways. By feeding BirA a chemically altered biotin — a ketone isostere in which the ureido nitrogens are replaced by carbons — researchers showed the enzyme will ligate an unnatural handle onto the AviTag, which can then be tagged with a fluorophore through ketone-specific chemistry, giving a two-step, bioorthogonal cell-surface labeling method.^[7] The lesson is that the ligase is tolerant enough to be repurposed, and that tolerance is what the proximity-labeling methods exploit next.

Proximity labeling: BioID and TurboID

One of the most influential modern uses of biotinylation turns the ligase’s reactive intermediate into a tool for mapping which proteins sit near one another inside a living cell. The idea, introduced as BioID in 2012, uses a mutant BirA (the R118G variant, “BirA*”) that cannot hold onto the activated biotinyl-5′-AMP it makes.^[12] Instead the reactive intermediate leaks out of the active site and biotinylates lysines on any protein within a few nanometres before it hydrolyses. Fuse that promiscuous ligase to a protein of interest, add biotin to the cells, and every protein that comes close enough gets a biotin tag; the tagged neighbourhood is then captured on streptavidin beads and identified by mass spectrometry.^[12] Because the tag is covalent and nearly irreversible, harsh lysis buffers can be used to solubilise membranes and chromatin without losing the labeled proteins.

The weakness of the original BioID was speed: it needed 18 to 24 hours of labeling, too slow to catch fast or dynamic events. In 2018 a group used yeast-display directed evolution to engineer faster ligases, TurboID and miniTurbo.^[13] Starting from a more active BirA point mutant and running 29 rounds of selection, they compressed the labeling window from 18 hours to about 10 minutes. TurboID is a 35-kilodalton enzyme carrying 15 mutations relative to wild-type BirA; miniTurbo is a smaller 28-kilodalton version with the N-terminal domain removed and 13 mutations, which gives lower background and a tighter labeling-time window at the cost of some activity.^[13] Crucially, TurboID was active enough to extend proximity labeling into whole organisms — flies and worms — where the older enzyme barely worked. The whole family rests on the same chemistry this guide began with: a reactive biotin species, attached to nearby lysines, read out through streptavidin capture.

What biotinylated peptides are used for

Biotinylation is a means, not an end, and the biotin–streptavidin pair shows up wherever something needs to be captured, immobilised or detected. The main families of use, all of them in-vitro laboratory techniques, are worth laying out because they explain why so many research peptides are offered in biotinylated form.

• Affinity capture and pull-downs. A biotinylated bait peptide is immobilised on streptavidin beads and used to fish binding partners out of a lysate, or to purify the tagged molecule itself. The near-irreversible bond means very little bait is lost during washing.^[1]
• Immobilisation on surfaces. Streptavidin-coated plates, chips and sensor surfaces present a uniform anchor; a biotinylated peptide binds in a defined orientation, which is why biotin capture is a standard way to load molecules onto surface-plasmon-resonance and biolayer-interferometry sensors for binding studies.
• Detection and reporting. Streptavidin conjugated to an enzyme (horseradish peroxidase, alkaline phosphatase) or a fluorophore turns a biotin tag into a signal. This is the basis of many ELISA, Western-blot, immunohistochemistry and flow-cytometry readouts, and multiple biotins per target multiply the signal.^[1]
• Multimer and complex assembly. Because streptavidin has four biotin sites, it can nucleate ordered assemblies — the classic example being MHC tetramers built from four biotinylated MHC–peptide complexes around one streptavidin, used to identify antigen-specific T cells.^[1]
• Interaction and neighbourhood mapping. The BioID and TurboID methods above use enzymatic biotinylation to label and then identify proteins near a target in living cells.^[12][13]

In each case the biotin is chosen as the tag precisely because it is small, chemically well-defined, and read out through a binding event that is effectively permanent under assay conditions. None of these uses describes administration to a person or animal; they are bench techniques for studying molecules.

How a biotinylated peptide is characterised

Adding biotin changes a peptide in small, precise ways that the analytical laboratory can see, which is how a biotinylated peptide is confirmed and how its quality is judged. The defining question is usually not just “is the biotin there?” but “how many, and where?”

The first tool is mass spectrometry. Biotin and its linker add a precise, known mass to the peptide, so a successfully biotinylated molecule appears at a predictably higher mass than its parent — and the number of added biotins can be read directly from the mass shift. Peptide mapping, in which the molecule is digested and the fragments are analysed, can confirm that the biotin sits on the intended residue rather than a stray one. The principles are the same as for any identity confirmation, covered in What Is Mass Spectrometry. The second tool is reversed-phase HPLC, the standard purity method: because biotin and its linker are modestly hydrophobic, a biotinylated peptide usually shifts its retention time relative to the unmodified sequence, and the method resolves the labeled product from unreacted starting material and from over-labeled by-products. A deeper treatment is in What Is HPLC.

A third, biotin-specific measurement is the degree of labeling — the average number of biotins per molecule. The classic method is the HABA assay: the dye HABA (2-(4′-hydroxyazobenzene)benzoic acid) binds avidin and turns a characteristic colour; biotin displaces it, and the fall in absorbance at 500 nm reports how much biotin is present, from which the biotin-to-protein ratio is calculated.^[1] These are ordinary tools of peptide quality control. Patriot Labs tests every batch of the peptides it sells for purity by HPLC and for identity by mass spectrometry through an independent US laboratory, with the certificate of analysis published on each product page; reading those documents is covered in Understanding Peptide COA Testing.

What this guide does not establish

This guide explains a chemistry and a set of laboratory techniques, not a use in any living subject, and the boundaries are worth stating plainly.

• Nothing here describes using any peptide, biotinylated or otherwise, in a person or animal. Biotinylation is described only as a bench method for capturing, immobilising and detecting molecules in vitro.
• Biotinylation changes handling, not biology. A biotin tag is attached so a molecule can be found and captured; it does not confer or imply any therapeutic property, and nothing here should be read as an effect in a living system.
• Named methods and reagents are cited to explain the chemistry. BioID, TurboID, the AviTag, specific reagent arm lengths and affinity figures are given to illustrate how the technique works, not as recommendations or protocols for any particular experiment.
• Specific figures are as reported in the cited sources. Dissociation constants, spacer-arm lengths, labeling times and molecular weights come from the references and are representative values, not universal constants; the right choices differ for every molecule and assay.
• “Research grade” is not a therapeutic grade, and nothing here describes Patriot Labs’ sourcing beyond the testing facts stated above.

Frequently asked questions

What is biotinylation?

Biotinylation is the covalent attachment of biotin, a small vitamin (vitamin B7, molecular weight 244.31 g/mol), to a peptide, protein, nucleic acid or other molecule. The point of the tag is capture and detection: biotin binds the proteins streptavidin and avidin with one of the strongest non-covalent affinities in nature, so a biotinylated molecule can be pulled down, immobilised or detected through that pairing. Biotin is small enough that it rarely disturbs the function of what it is attached to, and several biotins can be added to one molecule to amplify a signal.

Why does biotin bind streptavidin so tightly?

Streptavidin and avidin each have a deep, hydrophobic binding pocket, lined with tryptophan residues, that is an almost perfect complement to the biotin molecule. Once biotin is inside, a network of about eight hydrogen bonds and a mobile loop that closes over the pocket like a lid lock it in place. The result is a dissociation constant on the order of 10 to the minus 14 molar for streptavidin and about 10 to the minus 15 molar for avidin, far tighter than a typical antibody-antigen bond and resistant to heat, detergents, proteases and extremes of pH. It is non-covalent but approaches covalent strength.

What is the difference between chemical and enzymatic biotinylation?

Chemical biotinylation uses a reactive biotin reagent that couples to a chemical group on the target, most often the primary amines of lysine side chains, so it labels many sites with little control over where or how many. Enzymatic biotinylation uses the bacterial enzyme biotin ligase (BirA), which attaches a single biotin to one specific lysine inside a short recognition peptide such as the 15-residue AviTag, giving one biotin at one known position and a defined orientation. Chemical labeling is fast and needs no genetic engineering; enzymatic labeling is site-specific but requires the target to carry the tag sequence.

What does an NHS-ester biotin react with?

An NHS-ester (N-hydroxysuccinimide ester) biotin reacts with primary amines: the epsilon-amino group on lysine side chains and the alpha-amino group at a protein's N-terminus. It forms a stable amide bond and releases the NHS group as a leaving group. The reaction is run at slightly basic pH, roughly 7 to 9, because it is the deprotonated amine that reacts; it competes with hydrolysis of the ester by water, which has a half-life of only a few hours at pH 7 and a few minutes at pH 9, so amine-containing buffers such as Tris and glycine must be avoided.

How do you biotinylate a peptide at a single site?

The cleanest way with synthetic peptides is to build the biotin in during solid-phase synthesis or to attach it through a single uniquely reactive group. A common route is to include one cysteine in the sequence and use a thiol-reactive maleimide-biotin, which reacts about a thousand times faster with a free sulfhydryl than with an amine at neutral pH, so only that cysteine is labeled. Adding biotin (or biocytin, a biotin-lysine) to the N-terminus on the resin is another single-site route. For expressed proteins, the enzymatic AviTag/BirA system places one biotin on one lysine.

What is the AviTag?

The AviTag, also called the acceptor peptide, is a 15-amino-acid sequence (GLNDIFEAQKIEWHE) that is recognised by the enzyme biotin ligase BirA. When a protein is genetically fused to this tag and incubated with BirA, biotin and ATP, the enzyme attaches a single biotin to the lysine in the middle of the tag and nothing else. It was minimised from BirA's natural substrate through peptide-library screening, and it lets researchers biotinylate a protein at one defined position, in vitro or inside living cells, without modifying any other protein.

What is BioID and proximity labeling?

BioID is a method for mapping which proteins are near a protein of interest inside a living cell. The protein is fused to a promiscuous mutant of biotin ligase that releases the reactive biotin intermediate (biotinoyl-5'-AMP) instead of holding onto it, so biotin spills out and tags any protein that comes within a few nanometres. Those tagged proteins are then captured on streptavidin beads and identified by mass spectrometry. The original BioID enzyme needed 18 to 24 hours of labeling; an evolved version called TurboID cut that to about ten minutes and works in flies and worms.

Why are spacer arms used in biotinylation reagents?

The biotin binding pocket of streptavidin sits several angstroms below the protein's surface, so a biotin attached flush against a large molecule can be hard for streptavidin to reach. A spacer arm — a short linker between the reactive group and the biotin — holds the biotin out where it is accessible, which improves capture. Reagents are sold with different arm lengths, for example about 13.5 angstroms for a plain NHS-biotin, 22.4 angstroms for a long-chain (LC) version, and around 29 angstroms for a PEG4 version; polyethylene-glycol spacers also add water solubility.

What is the difference between avidin, streptavidin and NeutrAvidin?

All three are tetrameric proteins that each bind four biotins very tightly. Avidin comes from egg white, is glycosylated and strongly positively charged (high isoelectric point), which makes it bind biotin slightly harder but also stick non-specifically to many surfaces. Streptavidin comes from the bacterium Streptomyces avidinii, carries no sugar and has a near-neutral charge, so it gives much lower background. NeutrAvidin is avidin with its sugar chains removed, which lowers its charge and non-specific binding while keeping high biotin affinity, making it a middle option between the two.

Is biotinylation reversible, and how is it measured?

The biotin-streptavidin bond normally needs harsh conditions to break, such as 6 to 8 molar guanidinium chloride at low pH or boiling in detergent, which usually denatures the captured protein. For gentler release, researchers use a weaker-binding biotin analogue (desthiobiotin) that elutes with excess biotin, or a pH-switchable analogue (iminobiotin) that binds at alkaline pH and releases near pH 4. The degree of biotinylation — how many biotins per molecule — is commonly estimated with the HABA assay, in which biotin displaces a dye from avidin and the absorbance change at 500 nm is read.

References

1. Biotinylation — overview of chemical and enzymatic methods, reagent chemistries, biotin–streptavidin affinity and applications. Wikipedia. https://en.wikipedia.org/wiki/Biotinylation
2. Streptavidin — structure, biotin affinity (K_d ~10⁻¹⁴ M), comparison with avidin, and engineered variants. Wikipedia. https://en.wikipedia.org/wiki/Streptavidin
3. Avidin — egg-white origin, glycosylation, tetramer, and K_d ~10⁻¹⁵ M for free biotin. Wikipedia. https://en.wikipedia.org/wiki/Avidin
4. Green, N. M. (1963). Avidin. 1. The use of [¹⁴C]biotin for kinetic studies and for assay. Biochemical Journal, 89(3), 585–591. doi:10.1042/bj0890585
5. Beckett, D., Kovaleva, E., & Schatz, P. J. (1999). A minimal peptide substrate in biotin holoenzyme synthetase-catalyzed biotinylation. Protein Science, 8(4), 921–929. doi:10.1110/ps.8.4.921. PMID: 10211839
6. Schatz, P. J. (1993). Use of peptide libraries to map the substrate specificity of a peptide-modifying enzyme: a 13 residue consensus peptide specifies biotinylation in Escherichia coli. Bio/Technology, 11(10), 1138–1143. doi:10.1038/nbt1093-1138. PMID: 7764094
7. Chen, I., Howarth, M., Lin, W., & Ting, A. Y. (2005). Site-specific labeling of cell surface proteins with biophysical probes using biotin ligase. Nature Methods, 2(2), 99–104. doi:10.1038/nmeth735
8. Howarth, M., Chinnapen, D. J.-F., Gerrow, K., et al. (2006). A monovalent streptavidin with a single femtomolar biotin binding site. Nature Methods, 3(4), 267–273. doi:10.1038/nmeth861. PMID: 16554831
9. Weber, P. C., Ohlendorf, D. H., Wendoloski, J. J., & Salemme, F. R. (1989). Structural origins of high-affinity biotin binding to streptavidin. Science, 243(4887), 85–88. doi:10.1126/science.2911722. PMID: 2911722
10. Hendrickson, W. A., Pähler, A., Smith, J. L., et al. (1989). Crystal structure of core streptavidin determined from multiwavelength anomalous diffraction of synchrotron radiation. PNAS, 86(7), 2190–2194. doi:10.1073/pnas.86.7.2190. PMID: 2928324
11. Chivers, C. E., Crozat, E., Chu, C., et al. (2010). A streptavidin variant with slower biotin dissociation and increased mechanostability (traptavidin). Nature Methods, 7(5), 391–393. doi:10.1038/nmeth.1450. PMID: 20383133
12. Roux, K. J., Kim, D. I., Raida, M., & Burke, B. (2012). A promiscuous biotin ligase fusion protein identifies proximal and interacting proteins in mammalian cells (BioID). Journal of Cell Biology, 196(6), 801–810. doi:10.1083/jcb.201112098. PMID: 22412018
13. Branon, T. C., Bosch, J. A., Sanchez, A. D., et al. (2018). Efficient proximity labeling in living cells and organisms with TurboID. Nature Biotechnology, 36(9), 880–887. doi:10.1038/nbt.4201
14. Holmberg, A., Blomstergren, A., Nord, O., et al. (2005). The biotin–streptavidin interaction can be reversibly broken using water at elevated temperatures. Electrophoresis, 26(3), 501–510. doi:10.1002/elps.200410070. PMID: 15690449
15. Eakin, R. E., McKinley, W. A., & Williams, R. J. (1940). Egg-white injury in chicks and its relationship to a deficiency of vitamin H (biotin). Science, 92(2384), 224–225. doi:10.1126/science.92.2384.224. PMID: 17743857
16. Argaraña, C. E., Kuntz, I. D., Birken, S., Axel, R., & Cantor, C. R. (1986). Molecular cloning and nucleotide sequence of the streptavidin gene. Nucleic Acids Research, 14(4), 1871–1882. doi:10.1093/nar/14.4.1871. PMID: 3951999
17. Thermo Scientific. Avidin–Biotin Technical Handbook (reagent chemistries, spacer-arm lengths, sulfo- vs non-sulfo esters, HABA assay). https://tools.thermofisher.com/content/sfs/brochures/1601675-Avidin-Biotin-Handbook.pdf

For related reading, see What Is PEGylation? and What Is Lipidation? for two other ways a peptide is chemically modified, What Is Solid-Phase Peptide Synthesis? for how a biotin tag is built into a peptide on the resin, What Is Mass Spectrometry for confirming the added mass, and Understanding Peptide COA Testing for how identity and purity are shown.

All Patriot Labs products are sold strictly for in-vitro research and laboratory use only. Not for human or veterinary consumption. This guide is educational and describes the chemistry of biotinylation in general terms; it is not medical advice, does not describe how to use any product, and makes no therapeutic claim about any compound.