Protein Labeling, Crosslinking, and Modification Support—Getting Started

Protein interactions fundamentally can be characterized as stable or transient. Stable interactions are those associated with proteins that are purified as multi-subunit complexes and are best characterized by co-immunoprecipitation, pulldown or far-western methods. 

Transient interactions are on/off or temporary in nature and can be strong or weak, fast or slow. Such transient events can be captured by crosslinking or label transfer methods. 

The guides below offer a complete listing of available products:

• Crosslinking Reagents Technical Handbook
• Protein Interaction Technical Handbook
• Overview of Protein Labeling
• Overview of Crosslinking and Protein Modification

Crosslinking is the process of chemically joining two or more molecules by a covalent bond to form a novel complex having the combined properties of its individual components. 修飾は、元の分子の溶解性またはその他の特性を変えるための化学基の結合または切断を伴います。

"Labeling" generally refers to any form of crosslinking or modification whose purpose is to attach a chemical group (e.g., a fluorescent molecule) to aid in detection.

Protein crosslinking is used for many purposes, which include the following:

• Stabilize protein structure for analysis
• Capture and identify unknown protein interactors or interaction domains
• Conjugate an enzyme or tag to an antibody or other purified protein
• Immobilize antibodies or other proteins for assays or affinity purification
• Attach peptides to larger "carrier" proteins to facilitate handling/storage

In many situations, specialized protein modifications are needed to add molecular mass, increase solubility for storage, or create a new functional group that can be targeted in a subsequent reaction step.

Covalent modification and crosslinking of proteins depends on the availability of particular chemicals that are capable of reacting with the specific kinds of functional groups that exist in proteins. In addition, protein function and structure are either the direct focus of study or they must be preserved if a modified protein is to be useful in a technique. Therefore, the composition and structure of proteins, and the potential effects of modification reagents on protein structure and function, must be considered.

When selecting a crosslinker, it is important to consider the following factors:

• In vivo vs in vitro labeling: If labeling live cells, the crosslinker must be cell permeable unless you are crosslink proteins only on the membrane surface. If requiring a crosslinker that does not diffuse through the plasma membrane, use a branched chain crosslinker.
• Metabolic vs random labeling: We offer modified amino acids and other substrates that can be added to culture medium and incorporated into proteins or other biomolecules during protein synthesis, post-translational modification, replication or other synthetic pathways. 
• Reactive groups on the target: Do you know what reactive groups are available or are sites of interest on the proteins or other targets? Do you require a bifunctional crosslinker or trifunctional crosslinker? We offer crosslinkers that can react with amines, azides, carboxyls, carbonyls, hydroxyls and sulfhydryls in both homo- and heterobifunctional forms and trifunctional crosslinkers. When the reactive group on the target is unknown and linkage may be random, consider using a non-specific photoreactive crosslinker. The photoreactive crosslinkers can bind to primary amines, double bonds or insert into C-H and N-H sites. 
• Timing of the linkage: Spontaneous crosslinkers will bind upon contact with the appropriate reactive group on the protein or other molecules. The rate of reaction depends on the reaction conditions, the concentration of the target and crosslinker, and the rate of diffusion. If a target is expressed or active during a distinct cellular event or under the control of an inducer and you wish to label the protein at this specific event or induction, the photoreactive crosslinkers can be activated upon exposing the sample to light at the desired time and duration. Various modified amino acids and other biomolecules can be incorporated by live cells by adding the substrate to the culture media. You would want to control the timing of the addition of the substrate to the cells to coincide with a cellular event or treatment. 
• Solubility: Most in vitro crosslinking reactions are performed under mild physiological conditions. In some instances when a molecule is insoluble in aqueous buffers, some crosslinking reactions can be done in organic solvents such as acetonitrile, DMSO or DMF.
• Distance between molecules: We offer zero-length crosslinkers as well as crosslinkers with spacer arms of various lengths, from zero length to >100 angstroms. Longer spacer arms have greater flexibility and reduced steric hindrance.
• Cleavability: Do you require a crosslinker that is cleavable or reversible? A cleavable crosslinker is recommended if you need to dissociate the linked molecules to recover the individual components? Cleavable crosslinkers are often employed in ‘bait and prey’ analysis.
• Final analysis method: Are the crosslinked proteins/molecules destined for analysis by gel electrophoresis, western blotting, or mass spectrometry? If eventual analysis will be by mass spectrometry, we offer deuterated analogs of standard ‘light’ crosslinkers for improved identification of components involved in the protein interaction.

Protein crosslinkers are selected on the basis of their chemical reactivities (i.e., specificity for particular functional groups) and other chemical properties that affect their behavior in different applications:

• Chemical specificity refers to the reactive target(s) of the crosslinkers reactive ends. A general consideration is whether the reagent has the same or different reactive groups at either end (termed homobifunctional and heterobifunctional, respectively).
• Spacer arm length refers to the molecular span of a crosslinker (i.e., the distance between conjugated molecules). A related consideration is whether the arm is cleavable (i.e., whether the linkage can be reversed or broken when desired).
• Water-solubility and cell membrane permeability of a crosslinker affect whether it can permeate into cells and/or crosslink hydrophobic proteins within membranes. These properties are determined by the composition of the spacer arm and/or reactive group.
• Spontaneously reactive or photoreactive groups in a crosslinker affect whether it reacts as soon as it is added to a sample or can be activated at a specific time by exposure to UV light.

Crosslinkers can be classified as homobifunctional or heterobifunctional:

• Homobifunctional crosslinkers have identical reactive groups at either end of a spacer arm, and generally they must be used in one-step reaction procedures to randomly "fix" or polymerize molecules containing like functional groups. For example, adding an amine-to-amine crosslinker to a cell lysate will result in random conjugation of protein subunits, interacting proteins and any other polypeptides whose lysine side chains happen to be near each other in the solution. This is ideal for capturing a "snapshot" of all protein interactions but cannot provide the precision needed for other types of crosslinking applications.
• Heterobifunctional crosslinkers possess different reactive groups at either end. These reagents not only allow for single-step conjugation of molecules that have the respective target functional groups, but they also allow for sequential (two-step) conjugations that minimize undesirable polymerization or self-conjugation. In sequential procedures, heterobifunctional reagents are reacted with one protein using the most labile group of the crosslinker first. After removing excess non-reacted crosslinker, the modified first protein is added to a solution containing the second protein where reaction through the second reactive group of the crosslinker occurs.

SPB (Cat. No. 23013) is an amine- and photoreactive heterobifunctional crosslinking reagent. The NHS group will react with amines on the antibody while the psoralen group will bind to the oligo. The photoreactive psoralen group provides a much higher yield than phenyl azide-containing compounds, which makes this reagent a superior alternative to typical photoreactive NHS-ester crosslinkers. The psoralen tricyclic planar ring system intercalates into double-stranded, and to a lesser extent, single-stranded, DNA, RNA or oligonucleotides. 

Alternatively, if the oligo is modified with an amine group, you could use a homobifunctional crosslinker such as BS3 (Cat. No. 21580) or if the oligo is modified with thiol group, use a heterobifunctional crosslinker such as sulfo-SMCC (Cat. No. 22322). 

Our PMPI crosslinker (Cat No. 28100) is a maleimide-and-isocyanate crosslinker for attaching compounds to sulfhydryl groups after conjugating the linker to hydroxyl groups by a urethane (carbamate) bond. The first reaction is conjugating the hydroxyl group with the isocyanate. This will create a maleimide-activated molecule. 

You could then attach thiol groups on the protein using 2-Iminothiolane (Product No. 26101) or SATA (Product No. 26102). Remove excess reagent by dialysis or desalting and then react with the maleimide-activated molecule in the first step above.

A commonly used method for conjugating HRP to antibodies is by activation of the HRP using sodium meta-periodate (Cat no. 20504) to generate aldehydes. After the reaction, the HRP must be dialyzed to remove the meta-periodate; then the aldehydes on the HRP will react directly to the amine groups on the antibody. Alternatively, we offer activated HRP (Cat. No. 31488) as an alternative to avoid the periodate oxidation step to modify the HRP. 

There are three types of metabolic labeling reagents available:

• Click chemistry (azide-alkyne)
• Staudinger ligation (azide-phosphine)
• Photoreactive amino acids

The incorporation of azide-modified biomolecules permits eventual detection or crosslinking using alkyne or phosphine reagents. The incorporation of alkyne-modified biomolecules allows eventual detection or crosslinking using azide-modified reagents. The incorporation of either EdU or EU requires that the cells/tissues possess a pyrimidine salvage pathway. 

L-photo-leucine (Cat no. 22610) and L-photo-methionine (Cat no. 22615) are amino acid derivatives that possess diazirine rings for UV photo-crosslinking of proteins. When used in combination with specially formulated limiting media, these photo-activatable derivatives of leucine and methionine are treated like the naturally occurring amino acids by the protein synthesis machinery within the cell. As a result, they can be substituted for leucine or methionine in the primary sequence of proteins during synthesis. 

Incorporation of a metabolic labeling reagent is more selective than a photoreactive crosslinker in that the modified substrate is incorporated only during protein synthesis, post-translational modification, replication or other synthetic pathway. It may be incorporated into any biomolecule being synthesized during a specific cellular event or upon induction. Although you can control the timing and duration of crosslinking using a photoreactive crosslinker, all proteins, peptides or other biomolecules will be linked at the time of light exposure. 

Both the click and Staudinger reactions are chemoselective and bio-orthogonal, in that the reactive groups are specific to one another and are not found in biological systems. The use of these reagents results in minimal background and highly specific labeling or detection. 

Staudinger ligation involves the reaction of an azide with a phosphine to produce an aza-ylide intermediate that rearranges to form a stable amide bond and does not require copper. 

The click reaction links and azide with an alkyne and may require copper (classic ‘click’), lower copper (Click Plus) or may be done without copper (copperless click) using DIBO-alkyne reagents. 

We offer metabolic labeling reagents for incorporate that possess either an azide (e.g., AHA, 12-azidododecanoic acid) or alkyne (e.g. EdU, EU, HPG). The available detection and crosslinking reagents are also either azides (e.g., Invitrogen™ Alexa Fluor™ 488 azide, biotin azide), alkynes (e.g., Invitrogen™ Alexa Fluor™ 488 alkyne, Invitrogen™ Alexa Fluor™ 488 DIBO alkyne) or phosphines (e.g., Invitrogen™ DyLight™ 488-Phosphine, NHS-Phosphine).

Traditional spacer arms are composed of linear hydrocarbon chains that are not water soluble. Their hydrophobic property makes them suitable for crossing the plasma membrane, for intercellular and intramolecular crosslinking. The addition of a charged sulfonate or PEG makes the linear crosslinkers more water soluble, but may render them cell impermeant. 

Branched crosslinkers, usually with PEG, are water soluble but are unable to penetrate biological membranes due to the bulkiness of the branched chains. When a crosslinking application requires a crosslinker that must not cross the plasma membrane, branched crosslinkers are recommended. 

All wavelengths of light may cause photoactivation, but photoactivation is most efficient using UV wavelengths (250 to 370 nm). UV wavelengths do not penetrate through water, plastics or other materials very well. Most photoactivation reactions are performed with the tubes or flasks open and light shining directly upon the liquid or sample. The efficiency of photoactivation of a solution/sample is dependent upon the wavelength of light, the intensity of the light source, the distance of the light from the sample, exposure time, the volume and the depth of the solution, the extent of mixing (if any) during photoactivation, and the presence of other UV absorbing agents or other components that can block light. Solutions of larger volume or greater depth would require a longer exposure time relative to smaller volumes or shallower depths. Solutions containing components that can also absorb UV light or block light may require longer exposure times. Samples that contain solid material or are turbid are not as efficiently photoactivated relative to clear solutions. Viscous samples may require longer exposure or some method of consistent mixing to ensure all of the material is exposed to light. 

Although it is not required that you work in the dark when handling the reagents (pipetting, etc.) it is recommended to work under subdued artificial room light. If working in a room with sunlight, close the window shades. If working within a biosafety hood that has a sanitizing UV light, keep the reagents protected from light at all times. During any incubation with a photo-reactive reagent, it is best to cover culture dishes with foil or other covering to protect from exposure to light or prevent the possibility of accidental exposure to light.

We offer the following crosslinkers for protein to nucleic acids: formaldehyde (Cat. No. 28906), SPB (Cat. No. 23013), SDAD (Cat. No. 26169), sulfo-SDAD (Cat. No. 26175), and sulfo-SBED (Cat. No. 33034).

Formaldehyde can be used for DNA-DNA crosslinking as well as DNA-protein crosslinking. While it is good for crosslinking proteins that are in direct contact with DNA, formaldehyde cannot link proteins that may be bound in a complex with other proteins but are not in direct contact with DNA. EGS or DSG can crosslink proteins to other proteins that are in direct contact with DNA and crosslinked to DNA by formaldehyde.

SPB, Sulfo-SDAD, and Sulfo-SBED are all photoreactive crosslinkers. SPB is an NHS-ester and psoralen heterobifunctional crosslinker that conjugates primary amines on proteins to DNA via photo-activated intercalation of psoralen to pyrimidine bases. The psoralen tricyclic planar ring system intercalates into double-stranded, and to a lesser extent, single-stranded DNA and RNA. The photoreactive psoralen group provides more selective covalent binding to nucleic acids than either phenyl azide or diazerine linkers, which makes SPB the best choice. The diazerine on sulfo-SDAD and the phenyl azide on sulfo-SBED are not selective for nucleic acids and can crosslink other biomolecules in the same besides any protein-DNA interactions. Sulfo-SBED involves biotin transfer rather than actual linking.

Protein A as well as Protein A/G bind to antibodies via the Fc portion. You can use DSS crosslinker to crosslink the antibody to Protein A as we described it for Protein A/G in our kit (Cat No. 88805). Please note that with this chemical crosslinking, there is the potential to lose antigen binding sites, and thus, the IP yield may be a less than when performing the IP without crosslinking.

Please view our Tech Tip for a protocol on attaching a protein onto a gold surface. Another way to do this is to cleave hinge-region disulfides of the antibody with 2-MEA, which will produce free sulfhydryls to directly complex to the gold particle. Refer to Hermanson, Greg. (2013) Bioconjugate Techniques, Third Edition. New York, NY: Academic Press for detailed information. 

We offer Sufo-SBED (Cat. Nos. 33033 and 33034). 

We don't have any MS-labile crosslinkers. ‘MS labile’ means that the crosslinker will fragment or break down under HCD fragmentation. Most of our crosslinkers are stable in terms of covalently binding their target molecules but none of them have a fragmentation site for HCD. We do have crosslinkers that are cleavable using reducing agents or hydroxylamine.

The crosslinks formed by BS3 are stable to SDS/heat treatment as used for gels and blots. 

Amine-to-amine crosslinkers are a good choice for your application since amino groups are usually abundant on the surface of proteins. Our most popular amine-reactive homobifunctional crosslinker that is water-soluble is BS3 (Cat. No. 21585). This has a moderate-length (11.4 A) linker that is good for linking adjacent proteins. If you need the ability to cleave the linker with sulfhydryl reducing agents I would suggest DTSSP (Cat. No. 21578). 

If you think you might need an extra-long crosslinker then BS(PEG)5 (Cat. No. 21581) or BS(PEG)9 (Cat. No. 21582) are alternatives.

NHS-diazirine has not been tested for cross-linking in a ChIP application. EGS (Cat. No. 21565) and DSG (Cat. No. 20593) are not used by themselves in ChIP, but rather in combination with formaldehyde. While formaldehyde is good for crosslinking proteins that are in direct contact with DNA, it cannot trap proteins that may be bound in a complex with other proteins but are not in direct contact. Thus EGS or DSG can cross-link proteins to proteins that are in direct contact with DNA and are crosslinked to it by formaldehyde. See this reference for more information.

The working solution of BS3 can be dissolved (or diluted) in PBS, but concentrated solutions are not very soluble in solutions containing some salt. Use water, or, if buffer is required, no more than 10 mM sodium phosphate. Remember that stock solutions of BS3 (or any compound containing reactive NHS- or sulfo-NHS esters) cannot be stored as they will hydrolyze in aqueous solutions.

You can do the crosslinking on cells in plates, but use the very minimum amount of buffer that will cover the cells. Also before adding the BS3, be sure to carefully wash away any traces if serum and media components.

The density of reactive groups is reported on the lot-specific Certificate of Analysis. 

In many applications, it is necessary to maintain the native structure of the protein complex, so crosslinking is most often performed using near-physiologic conditions. Optimal crosslinker-to-protein molar ratios for reactions must be determined empirically, although product instructions for individual reagents generally contain guidelines and recommendations for common applications.

Depending on the application, the degree of conjugation is an important factor. For example, when preparing immunogen conjugates, a high degree of conjugation is desired to increase the immunogenicity of the antigen. However, when conjugating to an antibody or an enzyme, a low-to-moderate degree of conjugation may be optimal so that biological activity of the protein is retained.

The number of functional groups on the protein’s surface is also important to consider. If there are numerous target groups, a lower crosslinker-to-protein ratio can be used. For a limited number of potential targets, a higher crosslinker- to-protein ratio may be required. Furthermore, the number of components should be kept low or to a minimum because conjugates consisting of more than two components are difficult to analyze and provide less information on spatial arrangements of protein subunits.

Crosslinkers should be prepared as per the manufacturer’s instructions for the specific reagent. Hydrophobic crosslinkers are first dissolved in the appropriate solvent, such as DMF or DMSO, while more water-soluble crosslinkers (those with sulfonate groups or pegylated spacers) can be dissolved directly in water or aqueous buffer. 

The starting protein concentration or number of cells should be empirically determined for in vitro and in vivo crosslinking protocols, respectively. For in vitro crosslinking, the protein solution should be prepared in a nonreactive buffer, such as phosphate buffered saline (PBS) that has the proper pH for the specific crosslinker. For in vivo crosslinking applications, cells should be in the exponential phase of growth and at a subconfluent density during the crosslinking procedure. To avoid the possibility of culture media reacting with the crosslinker, the media can be replaced with PBS through a series of cell washes.

30 minutes is a good incubation time to start with, though multiple experiments can be performed concurrently to test other lengths of time to determine the optimal time of incubation with the specific crosslinker. Long incubation periods should generally be avoided, not only because it may cause formation of large, crosslinked protein aggregates, but also because the crosslinker may lose stability. In cases where extended incubation periods are required, though, fresh crosslinker can be added at specific time points throughout the procedure to maintain the proper molar ratio of reagent and maximize the formation of the target product. The formation of aggregates due to extensive crosslinking, though, should also be considered in determining the optimal reaction time.

To quantitate biotin, we offer two kits as follows:

• Biotin Quantitation Kit (Cat. No. 28005): With this kit, a solution containing the biotinylated protein is added to a mixture of HABA reagent (4’-hydroxyazobenzene-2-carboxylic acid and avidin. Because of its higher affinity for avidin, biotin displaces the HABA and the absorbance at 500nm decreases proportionally. 
• Fluorescence Biotin Quantitation kit (Cat. No. 46610): This microplate-based biotin assay is easy to perform by adding the supplied fluorescent reporter to the biotinylated samples and diluted biocytin standards. The avidin fluoresces when the weakly interacting HABA (4’-hydroxyazobenzene-2-carboxylic acid) is displaced by the biotin. The amount of biotin is determined by comparing the sample's fluorescence to the biocytin standard curve. This assay requires must less sample volume than the microplate colorimetric HABA assay and is much more sensitive

To determine the dye-to-protein ratio after fluorophore conjugation, absorbance readings of the protein:dye conjugate are taken and the molar ratio can then be calculated. Please go to this Tech Tip for more information.

Most labeled proteins can be stored in the same manner as the unmodified protein. The exact storage and stability is protein dependent. 

Simply stated, protein modification reagents are chemicals that block or expose reactive sites, alter native charges, inactivate function, or change functional groups to create targets for crosslinking and labeling. Three examples are sufficient to describe the types and purposes of modification reagents:

• Pegylation: Chemically attaching single- or branched-chain polyethylene glycol (PEG) groups to proteins is a form of labeling or modification that is primarily used to confer water-solubility and/or inert molecular mass to proteins. Forms of PEG that have been synthesized to contain reactive chemical groups comprise ready-to-use, activated reagents for pegylation.
• Block sulfhydryls: Protein sulfhydryls (side chain of cysteine) are important regulators of protein structure and function. Certain reagents are capable of reacting permanently or reversibly with sulfhydryl groups (e.g., NEM or MMTS, respectively). These reagents add a very small "cap" on the native sulfhydryl, enabling the activity of certain enzymes to be controlled for specific assay purposes.
• Convert amines to sulfhydryls: SATA and related reagents contain an amine-reactive group and a protected sulfhydryl group. By reacting the compound to a purified protein, the side chain of lysine residues can be modified to contain a sulfhydryl group for targeting with sulfhydryl-specific crosslinkers or immobilization chemistries. The method does not actually convert the amine into a sulfhydryl; rather it attaches a sulfhydryl-containing group to the primary amine. The effect is also to extend the length of the side chain by several angstroms.

The following functional groups are available for crosslinking/labeling:

• Primary amines (–NH2): This group exists at the N-terminus of each polypeptide chain (called the alpha-amine) and in the side chain of lysine (Lys, K) residues (called the epsilon-amine). Because of its positive charge at physiologic conditions, primary amines are usually outward-facing (i.e., on the outer surface) of proteins; thus, they are usually accessible for conjugation without denaturing protein structure.
• Carboxyls (–COOH): This group exists at the C-terminus of each polypeptide chain and in the side chains of aspartic acid (Asp, D) and glutamic acid (Glu, E). Like primary amines, carboxyls are usually on the surface of protein structure.
• Sulfhydryls (–SH): This group exists in the side chain of cysteine (Cys, C). Often, as part of a protein's secondary or tertiary structure, cysteines are joined together between their side chains via disulfide bonds (–S–S–). These must be reduced to sulfhydryls to make them available for crosslinking by most types of reactive groups.
• Carbonyls (–CHO): Ketone or aldehyde groups can be created in glycoproteins by oxidizing the polysaccharide post-translational modifications (glycosylation) with sodium meta-periodate.

A number of chemical reactive groups have been characterized and used to target the main kinds of protein functional groups. Some of the more popular groups are as follows:

• Carboxyl-to-amine reactive groups: Carbodiimide
• Amine reactive groups: NHS ester, Imidoester
• Sulfhydryl reactive groups: Maleimide, Haloacetyl, Pyridyldisulfide
• Aldehyde reactive groups: Hydrazide, Alkoxyamine
• Photoreactive (i.e., nonselective, random insertion) groups: Diazirine, Aryl Azide
• Hydroxyl reactive groups: Isocyanate

EDC and other carbodiimides are zero-length crosslinkers; they cause direct conjugation of carboxylates (–COOH) to primary amines (–NH2) without becoming part of the final crosslink (amide bond) between target molecules. EDC crosslinking reactions must be performed in conditions devoid of extraneous carboxyls and amines. Because peptides and proteins contain multiple carboxyls and amines, direct EDC-mediated crosslinking usually causes random polymerization of polypeptides. Nevertheless, this reaction chemistry is used widely in immobilization procedures (e.g., attaching proteins to a carboxylated surface) and in immunogen preparation (e.g., attaching a small peptide to a large carrier protein).

NHS esters are reactive groups formed by EDC-activation of carboxylate molecules. NHS ester-activated crosslinkers and labeling compounds react with primary amines in slightly alkaline conditions (pH 7.2-8.5) to yield stable amide bonds. The reaction releases N-hydroxysuccinimide (MW 115), which can be removed easily by dialysis or desalting. Primary amine buffers such as Tris or TBS are not compatible because they compete for reaction; however, in some procedures, it is useful to add Tris or glycine buffer at the end of a conjugation procedure to quench (stop) the reaction.

Sulfo-NHS esters are identical to NHS esters except that they contain a sulfonate (–SO3) group on the N-hydroxysuccinimide ring. This charged group has no effect on the reaction chemistry, but it does tend to increase the water-solubility of crosslinkers containing them. In addition, the charged group prevents sulfo-NHS crosslinkers from permeating cell membranes, enabling them to be used for cell surface crosslinking methods.

Imidoester crosslinkers react with primary amines to form amidine bonds. To ensure specificity for primary amines, imidoester reactions are best performed in amine-free, alkaline conditions (pH 10), such as with borate buffer.

Because the resulting amidine bond is protonated, the crosslink has a positive charge at physiological pH, much like the primary amine which it replaced. For this reason, imidoester crosslinkers have been used to study protein structure and molecular associations in membranes and to immobilize proteins onto solid-phase supports while preserving the isoelectric point (pI) of the native protein. However, the more stable and efficient NHS-ester crosslinkers have steadily replaced them in most applications.

Maleimide-activated crosslinkers and labeling reagents react specifically with sulfhydryl groups (–SH) at near neutral conditions (pH 6.5-7.5) to form stable thioether linkages. Disulfide bonds in protein structures (e.g., between cysteines) must be reduced to free thiols (sulfhydryls) to react with maleimide reagents. Extraneous thiols (most reducing agents) must be excluded from maleimide reaction buffers, because they will compete for coupling sites.

Short homobifunctional maleimide crosslinkers enable disulfide bridges in protein structures to be converted to permanent, irreducible linkages between cysteines. More commonly, the maleimide chemistry is used in combination with amine-reactive NHS-ester chemistry in the form of heterobifunctional crosslinkers that enable controlled, two-step conjugation of purified peptides and/or proteins.

Most haloacetyl crosslinkers contain an iodoacetyl or a bromoacetyl group. Haloacetyls react with sulfhydryl groups at physiologic to alkaline conditions (pH 7.2 to 9), resulting in stable thioether linkages. To limit free iodine generation, which has the potential to react with tyrosine, histidine and tryptophan residues, perform iodoacetyl reactions in the dark.

Pyridyl disulfides react with sulfhydryl groups over a broad pH range to form disulfide bonds. As such, conjugates prepared using these crosslinkers are cleavable with typical disulfide reducing agents, such as dithiothreitol (DTT).

During the reaction, a disulfide exchange occurs between the –SH group of the target molecule and the 2-pyridyldithiol group of the crosslinker. Pyridine-2-thione (MW 111; λmax 343nm) is released as a byproduct that can be monitored spectrophotometrically and removed from protein conjugates by dialysis or desalting.

Carbonyls (aldehydes and ketones) can be produced in glycoproteins and other polysaccharide-containing molecules by mild oxidation of certain sugar glycols using sodium meta-periodate. Hydrazide-activated crosslinkers and labeling compounds will then conjugate with these carbonyls at pH 5 to 7, resulting in formation of hydrazone bonds.

Hydrazide chemistry is useful for labeling, immobilizing or conjugating glycoproteins through glycosylation sites, which are often (as with most polyclonal antibodies) located at domains away from the key binding sites whose function one wishes to preserve.

Although not currently as popular or common as hydrazide reagents, alkoxyamine compounds conjugate to carbonyls (aldehydes and ketones) in much the same manner as hydrazides.

Photoreactive reagents are chemically inert compounds that become reactive when exposed to ultraviolet or visible light. Historically, aryl azides (also called phenylazides) have been the most popular photoreactive chemical group used in crosslinking and labeling reagents.

When an aryl azide compound is exposed to UV light, it forms a nitrene group that can initiate addition reactions with double bonds or insertion into C-H and N-H sites or can undergo ring expansion to react with a nucleophile (e.g., primary amine). Reactions can be performed in a variety of amine-free buffer conditions to conjugate proteins or even molecules devoid of the usual functional group "handles".

Photoreactive reagents are most often used as heterobifunctional crosslinkers to capture binding partner interactions. A purified bait protein is labeled with the crosslinker using the amine- or sulfhydryl-reactive end. Then this labeled protein is added to a lysate sample and allowed to bind its interactor. Finally, photo-activation with UV light initiates conjugation via the phenyl azide group.

Diazirines are a newer class of photo-activatable chemical groups that are being incorporated into crosslinking and labeling reagents. The diazirine (azipentanoate) moiety has better photostability than phenyl azide groups, and it is more easily and efficiently activated with long-wave UV light (330-370 nm).

Photo-activation of diazirine creates reactive carbene intermediates. Such intermediates can form covalent bonds through addition reactions with any amino acid side chain or peptide backbone at distances corresponding to the spacer arm lengths of the particular reagent. Diazirine-analogs of amino acids can be incorporated into protein structures by translation, enabling specific recombinant proteins to be activated as the crosslinker.

Many different molecules, including biotin, reporter enzymes, fluorophores and radioactive isotopes, can be attached to a target protein or nucleotide sequence. These labels are covalently attached and facilitate detection or purification of the labeled protein and/or its binding partners. さまざまなタイプの標識法がありますが、特定のアプリケーションによっては好ましい方法があります。 Therefore, the type of label and the labeling strategy used must be carefully considered and tailored for each application.

Biotin (vitamin H) is a useful label for protein detection, purification and immobilization because of its extraordinarily strong binding to avidin, streptavidin or NeutrAvidin™ Protein. Indeed, this interaction is one of the strongest non-covalent interactions between a protein and ligand. Additionally, biotin (244.3 Da) is considerably smaller than enzyme labels and is therefore less likely to interfere with normal protein function. Together, these features make avidin-biotin strategies ideal for many detection and immobilization applications. However, depending on the nature of the application, the very strong binding interaction can be problematic. In those situations, certain variants of avidin or derivatives of biotin are available, which allow soft-release (elution) binding or cleavable (reversible) labeling.

Biotinylation is the process of labeling proteins or nucleotides with biotin molecules and can be performed by enzymatic and chemical means. Chemical methods of biotinylation are most commonly used, and the biotinylation reagents used for this type of labeling share several basic features. They are composed of the biotinyl group, a spacer arm and a reactive group that is responsible for attachment to target functional groups on proteins. Variations in these three features account for the many varieties of available reagents and provide the specific properties needed for particular applications.

Active site probes are a class of chemical labeling reagents whose reactive groups are designed to specifically bind (label) particular enzyme active sites. Similar to traditional chemical labeling probes, active site probes contain a detectable tag (biotin/dye), a spacer arm, and a reactive group that is responsible for attachment to the active site of the target class of enzymes. Active site reactive groups are typically electrophilic compounds that covalently link to nucleophilic residues found in enzyme active sites. In cases where the active site reactive group does not covalently bond to the target enzyme, photo-reactive groups are incorporated into the linker region to facilitate attachment following specific binding. These probes can be used to selectively enrich, identify, and profile target enzyme classes across samples or assess the specificity and affinity of enzyme inhibitors. 

Active site probes have been developed to label different specific enzyme classes such as kinases, phosphatases, GTPases, serine hydrolases, cysteine proteases, metalloproteases, and cytochrome p450 enzymes. All active site probes can be used to determine inhibition of enzymes by small molecules, and some probes also preferentially react with only active enzymes, allowing for activity-based proteomic profiling (ABPP). Activity-based proteomic profiling is a powerful method to monitor protein activity versus traditional protein or RNA expression profiling techniques which only measure abundance.

Certain enzymes have properties that enable them to function as highly sensitive probes with a long shelf life and versatility for the detection of proteins in tissues, whole cells or lysates. Enzyme labels are considerably larger than biotin and require the addition of a substrate to generate a chromogenic, chemiluminescent or fluorescent signal that can be detected by different approaches. Enzyme labels are widely used because of their multiple types of signal output, signal amplification and the wide selection of enzyme-labeled products, especially antibodies.

Enzymes commonly used as labels include horseradish peroxidase (HRP), alkaline phosphatase (AP), glucose oxidase and β-galactosidase, and specific substrates are available for each enzyme. Indeed, multiple commercial substrates are available for HRP and AP that generate colorimetric, chemiluminescent or fluorescent signal outputs.

Fluorescent molecules, also called fluorophores or simply fluors, respond directly and distinctly to light and produce a detectable signal. Unlike enzymes or biotin, fluorescent labels do not require additional reagents for detection. This feature makes fluorophores extremely versatile and the new standard in detecting protein location and activation, identifying protein complex formation and conformational changes, and monitoring biological processes in vivo.

The vast selection of fluorophores today provides greater flexibility, variation and fluorophore performance for research applications than ever before. Fluorophores can be divided into three general groups, and each group of probes has distinct characteristics. These groups are as follows:

• Organic dyes—FITC, TRITC, Alexa Fluor™ dyes, DyLight™ Fluors
• Biological fluorophores—Green fluorescent protein (GFP), R-Phycoerythrin
• Qdot™ nanocrystals

Detection of fluorescent probes requires specialized equipment, including an excitation light source, filter set and a detector, that are found in fluorescence microscopes, fluorescence plate-readers, flow cytometers and cell sorters. This equipment enables the absolute quantitation of proteins based on fluorescence, which is a significant benefit to using fluorescent probes over other types of probes.