Hermanson G. Bioconjugate Techniques, Second Edition

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1000 27. Nucleic Acid and Oligonucleotide Modiﬁ cation and Conjugation

done using ﬂ uorescent probes to target particular regions within chromosomes. Called FISH

for ﬂ uorescent in situ hybridization, the technique is used extensively to identify marker chro-

mosomes or chromosomal rearrangements. Since many genomic sequences are repeated, usu-

ally occurring in multiple copies within isolated regions of the chromosome, the ﬂ uorescent

label on the DNA probe allows localization of targeted genes with high sensitivity. For a review

of FISH, see Meyne (1993).

Fluorescently labeled DNA probes also can be used in homogeneous assay systems to detect

and quantify target complementary sequences. The majority of these systems use a process of

energy transfer and ﬂ uorescent quenching to detect hybridization phenomena. The principle of

these assays involves the labeling of two binding components that can speciﬁ cally interact with

a target DNA. One or both of the labels may be a luminescent compound. The luminescent

quality of the ﬁ rst label may consist of a chemiluminescent probe that can be excited through

speciﬁ c chemical processes, producing light emission. Alternatively, the label may be a ﬂ uores-

cent probe which can absorb light of a particular wavelength and subsequently emit light at

another wavelength.

The second label also may be a ﬂ uorescent compound, but doesn ’t necessarily have to be.

As long as the second label can absorb the emission of the ﬁ rst label and modulate its signal,

binding events can be observed. Thus, the two labeled DNA probes interact with each other to

produce ﬂ uorescence modulation only after both have bound target DNA and are in enough

proximity to initiate energy transfer. Common labels utilized in such assay techniques include

the chemiluminescent probe, N -(4-aminobutyl)- N-ethylisoluminol, and reactive ﬂ uorescent

derivatives of ﬂ uorescein, rhodamine, and the cyanine dyes (Chapter 9). For a review of these

techniques, see Morrison (1992).

To prepare labeled DNA molecules for use in ﬂ uorescent assays, the oligo must be ﬁ rst deriv-

atized to contain a functional group. Any of the methods of Sections 2.1 and 2.2 (this chap-

ter) may be used to add an amine or sulfhydryl residue to speciﬁ c regions of the DNA polymer.

Once modiﬁ ed in this manner, the oligo may be further reacted with a ﬂ uorescent probe to cre-

ate the ﬁ nal derivative. However, many of the ﬂ uorescent quenching formats exclusively specify

either 3 - or 5 -labeled DNA molecules. This is because discrete modiﬁ cation at just one end of

the oligo assures that the label on a hybridized probe will be near enough to a second hybrid-

ized-and-labeled DNA molecule to effect the luminescent modulation necessary to make the

system viable. Multiple ﬂ uorescent labels on nucleotides within the DNA probe would not be

affected to the same degree by a second label attached to another oligo hybridized some dis-

tance away on the target strand. Therefore, use of the terminal transferase method of adding a

modiﬁ ed nucleoside triphosphate to the 3  end (Section 1) or 5 -phosphate modiﬁ cation using

a carbodiimide-mediated reaction (Section 2.1) work best for creating functionalized DNA

derivatives for ﬂ uorescent modulation techniques.

Another method of ﬂ uorescent detection of DNA or RNA targets involves the modiﬁ ca-

tion of a targeting oligo at both ends. In this approach, one end is modiﬁ ed with a ﬂ uorescent

molecule and the other end is modiﬁ ed with another label that can be a quencher of ﬂ uores-

cence or another ﬂ uorescent probe able to accept energy from the ﬁ rst label. These “molecular

beacons”, as they are called, contain terminal sequences that are able to hybridize the ends

together in solution in the absence of target. Thus, if a ﬂ uorescent molecule is at one end of the

molecular beacon and a quencher is at the other end, in solution without a target DNA present

the ﬂ uorescent signal would be completely quenched. In the presence of the target DNA, the

molecular beacon would hybridize to the target and open up the stem-and-loop structure of the

probe, thus forcing the ﬂ uorophore and quencher too far away from each other to cause ﬂ uo-

rescent quenching. The result is that speciﬁ c target binding of the molecular beacon causes the

generation of a ﬂ uorescent signal. The more ﬂ uorescence that occurs the more target is present

in solution. For a review of molecular beacon technology, see Bratu (2006).

The following sections describe two methods of coupling ﬂ uorescent labels to functional-

ized DNA probes. Other ﬂ uorophores or quenching molecules may be attached using similar

procedures with careful reference to the properties and reactivities of such labels as discussed

in Chapter 9.

Conjugation of Amine-Reactive Fluorescent Probes to Diamine-Modiﬁ ed DNA

DNA modiﬁ ed with a diamine compound to contain terminal primary amines may be coupled

with amine-reactive ﬂ uorescent labels. The most common ﬂ uorophores used for oligonucle-

otide labeling are the cyanine dyes and derivatives of ﬂ uorescein and rhodamine (Chapter 9).

However, any of the amine-reactive labels discussed throughout Chapter 9 are valid candidates

for DNA applications.

Some ﬂ uorescent probes are water-insoluble and must be dissolved in an organic solvent

prior to addition to an aqueous reaction medium containing the DNA to be labeled. Even

water-soluble ﬂ uorescent probes may be dissolved ﬁ rst in an organic solvent to permit easy

addition to an aqueous reaction medium without hydrolysis of the reactive group. Suitable sol-

vents are identiﬁ ed for each ﬂ uorophore, but mainly DMF or DMSO are used to prepare a

stock solution. Some protocols utilize acetone when labeling DNA. However, avoid the used

of DMSO for sulfonyl chloride compounds as this group reacts with the solvent. For oligo-

nucleotide labeling, the amount of solvent added to the reaction mixture should not exceed

more than 20 percent (although at least one protocol calls for a 50 percent acetone addition—

Nicolas et al ., 1992).

The following protocol is a generalized method for labeling amine-modiﬁ ed oligonucleotides

with a ﬂ uorescent probe, such as FITC. It is based on the method of Morrison (1992).

Protocol

1. Prepare 10 nmol of a diamine-modiﬁ ed DNA probe using the chemical methods dis-

cussed in Section 2.1 or through enzymatic derivatization using an amine-containing

nucleoside triphosphate (Section 1). Dissolve or buffer exchange the oligo into 1.0 ml of

a suitable coupling buffer for the type of amine-reactive ﬂ uorophore utilized (see recom-

mended reaction conditions for the particular ﬂ uorescent label in Chapter 9). For FITC,

the appropriate buffer condition for the oligo is 0.1 M sodium carbonate, pH 9.0.

2. Dissolve the ﬂ uorophore in DMF or another suitable solvent at a concentration of

0.01 M. For FITC, this translates into a concentration of 3.89 mg/ml.

3. Add 50 l of the FITC solution to the oligo solution and mix. For the use of NHS ester

or sulfonyl chloride ﬂ uorescent probes, add up to 150 l of the ﬂ uorophore solution to

the DNA.

4. React overnight at room temperature.

5. Remove excess ﬂ uorophore from the labeled oligo using gel ﬁ ltration on a desalting resin,

dialysis, or a centrifugal concentrator.

2. Chemical Modiﬁ cation of Nucleic Acids and Oligonucleotides 1001

1002 27. Nucleic Acid and Oligonucleotide Modiﬁ cation and Conjugation

Conjugation of Sulfhydryl-Reactive Fluorescent Probes to Sulfhydryl-Modiﬁ ed DNA

Fluorescent probes containing sulfhydryl-reactive groups can be coupled to DNA molecules

containing thiol modiﬁ cation sites. The chemical derivatization methods outlined in Section 2.2

(this chapter) may be used to thiolate the oligo for subsequent modiﬁ cation with a ﬂ uorophore.

Appropriate ﬂ uorescent compounds and their reaction conditions may be found in Chapter 9.

The protocol discussed in the previous section can be used as a general guide for labeling DNA

molecules.

1003

The study of protein interactions has become a vital research effort as a result of the

sequencing of the human genome and the genomes of other organisms. The next great challenge

beyond merely having knowledge of gene sequences is to understand the complex interplay of

the resultant protein molecules within cells as they bind, interact, and affect cellular processes.

From the many research papers that have appeared on this subject, it is becoming clear that

each protein molecule interacts with other proteins and molecules not in isolated obscurity, but

in a highly complex web of interactions, which can have far reaching effects on overall biologi-

cal function.

Protein interactions mediate virtually every cellular process. They are involved with tran-

scription, translation, transport, cell cycle control, the determination of cell type and function,

protein folding, post-translational modiﬁ cations, signal transduction, metabolism and energy

production, cell structure and motility, the formation of biological machines, cell and organism

defense, and apoptosis. Genetic mutations or damaging modiﬁ cations resulting in abnormal

protein interactions often are the root cause of many diseases, especially tumorigenesis.

The general types of protein–protein interactions that occur in cells include receptor–ligand,

enzyme–substrate, multimeric complex formations, structural scaffolds, and chaperones.

However, proteins interact with more targets than just other proteins. Protein interactions can

include protein–protein or protein–peptide, protein–DNA/RNA or protein–nucleic acid, protein–

glycan or protein–carbohydrate, protein–lipid or protein–membrane, and protein–small molecule

or protein–ligand. It is likely that every molecule within a cell has some kind of speciﬁ c interaction

with a protein.

The consequences of protein interactions in effecting cellular biochemistry include the syn-

thesis, destruction, or recycling of biomolecules; the supply of energy for cellular processes;

the generation of chemical signals; activation or inhibition of proteins and enzymes; changes

in protein conformation and structure; the creation of new binding sites or active centers in

proteins; the motility of cells, tissues, and organisms; transport of molecules within cells or

into/out of cells; and the formation of cellular structures and compartments.

Protein interactions can be described in relative ways related to the strength of the binding

that takes place between the two molecules and the time that the interaction lasts. Any inter-

acting molecules can be characterized as having either (1) high afﬁ nity (strong) or low afﬁ nity

(weak) binding and (2) stable (long lasting) or transient (short-lived) binding. Thus, a given

Bioconjugation in the Study of Protein Interactions

protein interaction may be described in one of four possible ways: strong and stable, strong

and transient, weak and stable, or weak and transient. Interactions of the weak and stable type

may at ﬁ rst seem like an oxymoron, but often multiple weak interactions can take place simul-

taneously with a target and the combined “avidity” makes the resultant complex stable.

Of the large number of protein interactions that take place in cells, perhaps the vast major-

ity may be described as transient. Most proteins that modify other molecules do so very rapidly

and so interact only brieﬂ y with their substrates or binding partners (i.e., enzymes). In addition,

since proteins within cells are highly compartmentalized, the afﬁ nity of most interactions doesn ’t

have to be very great, because each potential binding partner is within short diffusion distances

and the relative concentration of molecules within these small volumes is high.

Of course, the designations of strong, weak, stable, or transient are all subjective terms. They

mainly result from the outcome of afﬁ nity capture experiments of binding partners on insolu-

ble supports or the analytical determination of the kinetic parameters of speciﬁ c binding inter-

actions. In general, if the afﬁ nity constant of a protein interaction is strong enough to allow

a binding partner to be captured and puriﬁ ed using an immobilized afﬁ nity ligand, then the

interaction can be described as being reasonably strong. This usually correlates to an afﬁ nity

constant of 10

/M. Conversely, interactions having afﬁ nity constants of 10

/M are often too

weak to survive the washing steps needed to isolate the interacting protein on a solid phase

afﬁ nity support. Quantitative measurement of the afﬁ nity constant between interacting proteins

and the half-life of the interaction may be done using surface plasmon resonance (SPR) tech-

niques (Homola et al., 1995; Homola et al., 1999).

The vast network of protein–protein interactions that have been deciphered in recent years

has grown to include literally thousands of proteins in a sometimes-chaotic dance of complex-

ity. Using the two-hybrid method for instance, which involves the use of split transcription fac-

tor fusion proteins that allow detection of interacting proteins by activation of reporter gene

expression, many putative protein–protein interaction partners have been identiﬁ ed in yeast

(Fields and Song, 1989; Chien et al., 1991; Criekinge and Beyaert, 1999). A map of these inter-

actions looks a lot like a picture of the myriad nodes of addresses on a vast network of com-

puters, such as the interconnections that make up the Internet (Jeong et al., 2001). Although

most of the proteins in an organism have only a few partners that interact with them, major

hubs in protein “interactomes” can have up to 10–20 links with other proteins, indicating that

these key proteins are critical players in cell vitality Figure 28.1 .

Through the growing knowledge of protein–protein interactions and their corresponding gene

sequences major interaction domains on protein surfaces are being identiﬁ ed. These relatively

conserved amino acid sequences create structural motifs that are designed to bind with certain

targeted sequences in other proteins or molecules (Pawson and Nash, 2003; Ingham et al., 2005).

Using this information, many common binding regions on proteins can be identiﬁ ed just through

their gene sequences.

However, it is much more difﬁ cult to characterize the interactions of proteins with no

known interaction domains. The traditional “lock and key ” approach to conceptualizing

binding pairs is far too simplistic to allow easy visual identiﬁ cation of interacting surfaces on

the complex three-dimensional space making up the topography of protein molecules. Even

in those instances where protein interacting partners have been crystallized together and their

three-dimensional structures determined, it ’s obvious from the molecular models that it would

be difﬁ cult or impossible to identify visually the site of interaction without having such struc-

tural data in place beforehand. For this reason, experimental schemes are needed that are more

1004 28. Bioconjugation in the Study of Protein Interactions

elaborate than just knowledge of genetic sequences or interaction domains to characterize the

majority of speciﬁ c interactions proteins undergo. Even when two-hybrid studies indicate the

probability of a protein interaction occurring, it still doesn ’t provide information on the nature

of the interaction, the binding sites, or its function.

The techniques developed to study protein interactions can be divided into a number of major

categories (Table 31.1), including bioconjugation, protein interaction mapping, afﬁ nity capture,

two-hybrid techniques, protein probing, and instrumental analysis (i.e., NMR, crystallography,

mass spectrometry, and surface plasmon resonance). Many of these methods are dependent on the

use of an initial bioconjugation step to discern key information on protein interaction partners.

Many of the methods developed to study protein interactions use the bait/prey model to

detect interacting partners (Phizicky and Fields, 1995; Archakov et al., 2003 Piehler, 2005).

The bait protein is a puriﬁ ed protein (often recombinant) that is used to lure and capture a

putative interacting protein or biomolecule. The bait protein may be immobilized to a solid

phase for afﬁ nity separations or be used in solution. It also may be fusion tagged (i.e., GST or

6 His) or labeled with a detectable molecule, such as a ﬂ uorescent probe. It often is the case

Figure 28.1 A small segment of the yeast interactome. The spheres represent proteins and the interconnecting

lines are identiﬁ ed protein interactions. Many proteins are seen to interact with one or two other proteins, but

some can have over a dozen other interacting partners.

28. Bioconjugation in the Study of Protein Interactions 1005

1006 28. Bioconjugation in the Study of Protein Interactions

that there exists an antibody speciﬁ c for the bait protein to use for detection or in recovery of

the interacting complexes.

The prey is a protein, protein complex, or other biomolecule that interacts with the bait

protein. It can be captured by its speciﬁ c afﬁ nity interaction with the bait. Since many pro-

tein interactions may involve low afﬁ nity binding events or are transient, the use of chemical

crosslinking techniques can greatly facilitate analysis of an interacting prey protein. In fact, the

use of bioconjugation to “ﬁ x ” interacting proteins is a powerful route to capturing weak afﬁ n-

ity or transiently interacting molecules, as the crosslinked complexes can be isolated and ana-

lyzed without loss of some components.

The following sections describe the use of bioconjugation reagents for the study of protein

interactions. These reagents and techniques can be used to crosslink and capture interacting pro-

teins, to investigate the binding sites involved with interactions, and to identify which peptide

regions or amino acids participate in the binding event. In addition to the bioconjugation rea-

gents described in this section, the reader also is referred to Chapter 16, entitled Mass Tags and

Isotope Tags, wherein some of those reagents also can be used to investigate interacting proteins.

1. Homobifunctional Crosslinking Agents

One of the ﬁ rst applications of bioconjugate techniques was the use of simple homobifunctional

compounds (see Chapter 4) to capture interacting proteins in biological samples. As the name

of this type of reagent implies, these compounds have reactive groups on each end of a spacer

arm that are identical and react with the same type of functionality on different proteins. For

instance, early development of crosslinking compounds resulted in the creation of homobifunc-

tional amine-reactive reagents using either imidoesters or NHS esters as the reactive groups.

Both of these reagent types could be reacted with a complex protein sample mixture to result in

efﬁ cient conjugation of proteins through their available amine groups. The reactive groups on

these compounds survive long enough in a physiological pH environment to effectively capture

and covalently link proteins in proximity to one another. In this way, any proteins undergoing

speciﬁ c biological interactions can be “ﬁ xed ” by chemical conjugation and thus link together

permanently interacting complexes for analysis.

However, as one might expect, this strategy is more or less a crude “shotgun” approach,

wherein every protein in the sample has the potential to be crosslinked with other proteins

regardless of whether they are undergoing speciﬁ c interactions or not. The use of homobifunc-

tional reagents to study protein interactions therefore may result in high noise levels or many

false positives, because of their non-selective crosslinking characteristics. The end result often

makes it difﬁ cult to identify conjugated proteins that are speciﬁ cally interacting from those pro-

teins crosslinked just due to random collisions.

However, even acknowledging their disadvantages, homobifunctional crosslinkers have

been used successfully to investigate many protein–protein interactions, within cells and within

lysates or protein solutions. The key to capturing true interacting proteins while limiting the

degree of nonspeciﬁ c conjugation is to optimize the amount of crosslinker at the lowest pos-

sible concentration. Using too high a concentration will extensively conjugate all proteins, even

those that are not speciﬁ cally interacting at the moment. Using too low a concentration will not

conjugate effectively even interacting proteins. Therefore, optimization typically is needed to

determine the best concentration of homobifunctional crosslinker for a particular application.

1.1. DSS and BS

Two crosslinking agents that have been used extensively to study protein interactions are disuc-

cinimidyl suberate (DSS) and bis-sulfosuccinimidyl suberate (BS

) (see Chapter 4, Section 1.2).

Both reagents contain an 8-carbon spacer arm built from suberate core and have reactive esters at

each end that couple with amines to form amide bonds. They differ, however, in the fact that DSS

contains NHS esters and BS

contains negatively charged sulfo-NHS esters. BS

therefore is water-

soluble and will not penetrate cell membranes, whereas DSS is hydrophobic, water-insoluble, and

is membrane permeable. For this reason, BS

can be used to study cell-surface–protein interactions

(Friedrichson and Kurzchalia, 1998; Simons et al., 1999), while DSS can be used to study intra-

cellular protein interactions (Ishmael et al., 2005). The reaction of DSS to capture and crosslink

interacting proteins is shown in Figure 28.2 .

The following protocol represents a generalized strategy for crosslinking interacting proteins

using either DSS or BS

. The buffer conditions and reagent amounts are gleamed from published

procedures, but the exact quantities should be optimized for each protein interaction studied.

Protocol

1. Suspend cells at  25  10

cells/ml in PBS (pH 8.0).

2. Wash cells 3 times with ice-cold PBS (pH 8.0) to remove amine-containing culture media

and extracellular proteins from the cells.

3. For cell-surface interaction studies, add ligands to the cells and incubate for 1 hour at

4°C.

4. Dissolve DSS or BS

in dry DMSO at a concentration of 25 mM. Note: BS

may be added

directly to PBS buffer or dissolved as a stock solution in DMSO.

5. Add an aliquot of the DSS or BS

solution to the reaction medium to obtain a ﬁ nal con-

centration of 0.5–5 mM. Note: Simons et al. (1999) successfully used a concentration of

0.5 mM BS

with Madin-Darby canine kidney (MDCK) cells permanently expressing a

GPI-anchored form of growth hormone decay accelerating factor (GH-DAF) to crosslink

the protein interaction complexes on the cell surfaces.

Figure 28.2 DSS can capture protein interacting partners through amide bond crosslinks.

1. Homobifunctional Crosslinking Agents 1007

1008 28. Bioconjugation in the Study of Protein Interactions

6. Incubate the reaction mixture for 30 minutes at room temperature. To reduce active

internalization of BS

into cells, this incubation may be performed at 4°C.

7. Quench the reaction by adding an aliquot of 1 M Tris, pH 7.5, to give a ﬁ nal concentra-

tion of 10–20 mM.

8. Incubate the quenching reaction for 15 minutes at room temperature.

9. Lyse cells and analyze the protein interactions by electrophoresis, Western blotting, and

mass spec.

1.2. Heavy Atom, Deuterated Crosslinking Agents

Another approach to the study of protein interactions using homobifunctional crosslinkers uses

the incorporation of isotopes, like deuterium, into the reagent structure to produce a heavy

atom analog suitable for mass spec analysis. In this technique, the normal or light H (hydro-

gen) version of a crosslinker is used in an equal molar ratio to a heavy D (deuterium) version

to capture interacting proteins through covalent conjugation. The heavy and light analogs are

reacted with a sample at the same time, so that each form of the reagent will have an equal

chance of reacting with an interaction complex. Subsequent proteolytic digestion of the sample

(i.e., with trypsin) creates peptide fragments, some of which will contain covalent crosslinks

from the heavy or light crosslinkers. In addition, these crosslinked peptides will have an equal

chance of having either a light atom linker or a heavy atom linker holding them together.

Mass spec analysis of the peptide fragments formed by this process yields pairs of MS peaks

differing only by the mass change caused by the substitution of deuterium atoms for hydrogen

atoms in half of the crosslinks. Thus, searching for MS peaks in the data that differ by the

number of deuterium substitutions immediately will identify peptides from the interacting pro-

teins that have been captured by the crosslinking process.

Using this approach, homobifunctional crosslinking agents containing sulfo-NHS esters have

been developed based on the core structures of glutaric and suberic acids. The suberate-based

crosslinker also is known as BS

and has been described previously (Section 1.1, this chapter).

These two standard amine-reactive reagents then are modiﬁ ed at two carbons of their respec-

tive cross-bridge structures to contain two pairs of deuterium atoms, increasing their molecular

mass by exactly 4 from the normal hydrogen atom analogs.

These heavy atom reagent pairs are termed BS

G-d

(bis[sulfosuccinimidyl] 2,2,4,4

glutarate-d

) and BS

-d

(bis[sulfosuccinimidyl] 2,2,7,7 suberate-d

), and their light atom ana-

logs are called BS

G-d

(bis[sulfosuccinimidyl] glutarate-d

) and BS

-d

(bis[sulfosuccinimidyl]

suberate-d

) (Thermo Fisher) ( Figure 28.3 ). The sulfo-NHS esters of these four compounds all

react equally well with amine groups on proteins to form amide bonds. Therefore, there are

virtually no differences in properties or reactivities between the heavy and light versions of

these reagents.

The conjugation of interacting proteins with heavy/light crosslinkers potentially can result

in a number of derivatives having unique atomic mass units (amu) observed by mass spec. For

instance, any of these crosslinkers might react with amines on interacting proteins at both ends

to form amide bonds. Alternatively, one end of these homobifunctional compounds may react

with a protein to form an amide bond while the other end hydrolyzes, resulting in loss of the

sulfo-NHS group to form a carboxylate. Thus, the heavy/light crosslinking pairs may have two

potential amu results when peptides are analyzed by mass spec. Table 31.2 shows the potential

amu contributions for each of the possible products formed by the reactions of the heavy/light

crosslinkers with proteins.

This technique has been described as a general method of studying protein–protein interac-

tions as well as a method for investigating the three-dimensional structure of individual proteins

(Muller et al., 2001; Back et al., 2003; Dihazi and Sinz, 2003; Sinz, 2003; Sinz, 2006). It also has

been used for the study of the interactions of cytochrome C and ribonuclease A (Pearson et al.,

2002), to investigate the interaction of calmodulin with a speciﬁ c peptide binder (Kalkhof et al.,

2005a; Schmidt et al., 2005), and for probing laminin self-interaction (Kalkhof et al., 2005b).

In practice, sample concentrations typically are kept in the micromolar range to limit the

widespread conjugation of molecules not speciﬁ cally interacting or to limit intermolecular con-

jugation when studying the structure of a single protein. Therefore, most protein concentra-

tions will be in the microgram/ml range prior to the addition of crosslinkers. Even in cases

wherein a single puriﬁ ed protein is crosslinked to study its three-dimensional structure, limit-

ing the amount of crosslinker in the reaction mixture will help to limit polymerization of the

protein. However, successful results have been obtained by this method using up to a 200-fold

excess of crosslinker over the concentration of protein in solution.

The following protocol is based on the published methods as well as the speciﬁ c instructions

provided with the heavy atom crosslinkers from Thermo Fisher Scientiﬁ c. For new applications,

the amount of crosslinkers added to the sample will have to be optimized to obtain useful

information about the interactions.

Figure 28.3 The homobifunctional crosslinkers BS

G and BS

can be used to capture protein interactions

through amide bond formation. The deuterium-labeled analogs of these reagents can be used to differentiate

samples by mass spec.

1. Homobifunctional Crosslinking Agents 1009