951Bioconjugate Techniques, Third Edition.DOI: © 2013 Elsevier Inc. All rights reserved.http://dx.doi.org/10.1016/B978-0-12-382239-0.00022-4

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Enzymes are widely used in bioconjugate chemistry as detection components in assay systems. The cata-lytic activity of an enzyme can be used to turn substrate molecules into chromogenic, fluorescent, or chemilumi-nescent products, which are easily detectable or quan-tifiable by imaging, microscopy, or spectroscopy. If an enzyme is conjugated to a targeting molecule specific for some analyte of interest, then an assay system can be constructed to localize or measure the analyte. The most common targeting molecule is an antibody hav-ing antigen-binding specificity for the substance to be measured. An enzyme conjugated to such an antibody can be used to visualize the presence of antigen. Due to the advantages of this simple concept, enzyme-linked immunoadsorbent assays (ELISAs) have become the most important type of immunoassay system available.

The rapid turnover rate of some enzymes allows ELISAs to be designed that far surpass the sensitivity of radiolabeling techniques. In addition, substrates can be chosen to produce soluble products that can be accu-rately quantified by their absorbance or fluorescence. Alternatively, substrates are available that form insolu-ble, highly colored precipitates, excellent for localizing antigens in blots, cells, or tissue sections. The flexibility of enzyme-based assay systems makes the chemistry of enzyme conjugation one of the most important applica-tion areas in bioconjugate techniques.

In addition, the immobilization of enzymes for use in catalytic transformations (called immobilized reac-tors) has also become an important field in the use of these proteins. Specialized immobilized reactors are being used to cleave or modify biological molecules, to synthesize complex organic compounds, to pro-duce food products, and for the production of bioen-ergy molecules from biomass feedstock. See Chapter 1 for a review of immobilized reactor applications and Chapter  15 for the activation and coupling methods that can be used to covalently attach enzymes to vari-ous types of insoluble support materials.

The following sections briefly describe the princi-pal enzymes used for conjugation with other protein

molecules, particularly in the design of ELISA and other immunoassay systems.

1. PROPERTIES OF COMMON ENZYMES

1.1. Horseradish Peroxidase (HRP)

HRP (donor:hydrogen peroxide oxidoreductase; EC 1.11.1.7), derived from horseradish roots, is a enzyme of molecular weight 40,000 that can catalyze the reaction of hydrogen peroxide with certain organic, electron-donating substrates to yield highly colored products (Figure 22.1). The reaction of HRP with its fundamen-tal substrate, H2O2, forms a stable intermediate that can dissociate in the presence of a suitable electron donor, oxidizing the donor and potentially creating a color change. The donor can consist of oxidizable molecules like ascorbate, cytochrome c, ferrocyanide, or the leuco forms of many dyes. A large variety of

C H A P T E R

Enzyme Modification and Conjugation

FIGURE 22.1 Horseradish peroxidase shown as the ribbon struc-ture with the heme ring in its active center and two bound calcium ions. The molecular model is based on structure 1H58 in the RCSB Protein Databank by Berglund et al. (2002).

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electron-donating dye substrates are commercially available for use as HRP detection reagents. Some of them can be used to form soluble colored products for use in spectrophotometric detection systems, while other substrates form insoluble products that are espe-cially appropriate for staining techniques. In addition, substrates are available that create fluorescent or che-miluminescent products upon oxidation with HRP. The chemiluminescent substrates are among the most sensi-tive of all detection reagents, facilitating the detection of as little as attogram quantities of many targeted ana-lytes. The pH optimum for HRP is 7.0 although particu-lar substrate detection reactions may be performed at pH values slightly different from neutrality.

The use of antibody–HRP or streptavidin–HRP con-jugates in peroxidase-catalyzed enhanced chemilumi-nescent assays can result in one of the most sensitive detection methods for assaying targeted analytes in ELISA and western blotting applications. The reaction cascade that occurs during HRP catalysis can be dra-matically improved by the addition of various enhancer molecules, which create oxidized intermediates leading to the oxidation and light emission of a chemilumines-cent substrate such as luminol. Vdovenko et  al. (2012) analyzed this reaction using a multi-factorial design of experiments (DOE) approach to identify the best com-bination and concentrations of H2O2, luminol, and two different enhancer compounds (3-(10′-phenothiazinyl)propane-1-sulfonate and 4-morpholinopyridine. This combination at an optimized concentration resulted in the best signal-to-noise ratio and the longest chemilu-minescent emission.

HRP is a hemoprotein containing photohemin IX as its prosthetic group. The presence of the heme structure gives the enzyme its characteristic color and maximal absorptivity at 403 nm. The ratio of its absorbance in solution at 403 nm to its absorbance at 275 nm, called the RZ or Reinheitzahl ratio, can be used to approximate the purity of the enzyme. However, at least seven iso-enzymes exist for HRP (Shannon et al., 1966; Kay et al., 1967; Strickland et  al., 1968), and their RZ values vary from 2.50 to 4.19. Thus, unless the RZ ratio is precisely known or determined for the particular isoenzyme of HRP utilized in the preparation of an antibody–enzyme conjugate, subsequent measurement after crosslinking would yield questionable results in the determination of the amount of HRP present in the conjugate.

HRP is a glycoprotein that contains significant amounts of carbohydrate. Its polysaccharide chains are often used in crosslinking reactions to couple the enzyme to targeting molecules. Mild oxidation of its associated glycan sugar residues with sodium periodate generates reactive aldehyde groups that can be used for conjugation to amine-containing molecules. Reductive amination of oxidized HRP to antibody molecules in

the presence of sodium cyanoborohydride is perhaps the simplest method of preparing highly active con-jugates with this enzyme (Chapter  4, Section 1.4, and Chapter 20, Section 1.3).

Other methods of HRP conjugation include the use of the homobifunctional reagent glutaraldehyde (Chapter  5, Section 6.2, and Chapter  15, Section 2.1) and the heterobifunctional crosslinker, SMCC (succin-imidyl-4-(N-maleimidomethyl)cyclohexane-1-carbox-ylate) (Chapter  6, Section 1.3). Using glutaraldehyde, a two-step protocol is usually employed to try to limit the extent of oligomer formation. Even using the most highly controlled reactions, however, this method often causes unacceptable amounts of precipitated conjugate. Despite this disadvantage, glutaraldehyde conjugation is still routinely used, especially in the preparation of some antibody–enzyme reagents that go into established diagnostic assays. The use of the N-hydroxysuccinimide (NHS) ester–maleimide cross-linker, SMCC, provides far better control over the con-jugation process. SMCC is usually reacted first with HRP to create a derivative containing sulfhydryl-reac-tive maleimide groups. HRP activation of the native enzyme should result in the modification of a maximum of about two amine groups on the protein, because HRP only contains two lysines. An increase in the activa-tion level can be realized if the enzyme first is modified with ethylenediamine (EDA) using the carbodiimide EDC according to the methods described in Chapter 19 for the production of cationized bovine serum albumin (cBSA). The EDA-modified HRP is also more stable than the unmodified version, so cationization may have ben-efits in the retention of enzyme activity. The maleimide-activated enzyme can be purified and freeze-dried, providing a ready source of modified HRP to react with a sulfhydryl-containing antibody. Several preactivated forms of this enzyme are available from Thermo Fisher.

The size of HRP is an advantage in preparing anti-body–enzyme conjugates, since the overall complex size also can be designed to be small. Relatively low-molecular-weight conjugates are able to penetrate cel-lular structures better than large, polymeric complexes. This is why HRP conjugates are often the best choice for immunohistochemical (IHC) and immunocytochemi-cal staining techniques. Small conjugate size means greater accessibility to antigenic structures within tissue sections.

Another distinctive advantage of HRP is its robust nature and stability, especially under the conditions employed for crosslinking. HRP is stable for years in a freeze-dried state, and the purified enzyme can be stored in solution at 4°C for many months with-out significant loss of activity. The enzyme also retains excellent activity after being modified with a conju-gation reagent or after being periodate-oxidized to

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form aldehyde groups on its polysaccharide chains. Depending on the methods used for crosslinking, HRP conjugates can be constructed to have a high ratio of enzyme to antibody or a low ratio—both retaining high specific activity.

The disadvantages associated with HRP are sev-eral. The enzyme only contains two available primary ε-amine groups—extraordinarily low for most pro-teins—thus limiting its ability to be activated with amine-reactive heterobifunctionals. HRP is sensitive to the presence of many antibacterial agents, especially azide. It is also reversibly inhibited by cyanide and sulfide (Theorell, 1951). Finally, while the enzymatic activity of HRP is extremely high, its useful life span or practical substrate development time is somewhat lim-ited. After about an hour of substrate turnover, in some situations its activity can be decreased severely.

Nevertheless, HRP is by far the most popular enzyme used in antibody–enzyme conjugates. One sur-vey of enzyme use stated that HRP is incorporated in about 80% of all antibody conjugates, most of them uti-lized in diagnostic assay systems.

1.2. Alkaline Phosphatase

Alkaline phosphatases [AP, orthophosphoric mono-ester phosphorylase (alkaline optimum); EC 3.1.3.1] represent a large family of almost ubiquitous isoen-zymes found in organisms from bacteria to animals (Figure 22.2). In mammals, there are two forms of AP, one form present in a variety of tissues and another form found only in the intestines. They share common attributes in that the phosphatase activity is optimal at pH 8 to 10, is activated by the presence of divalent cat-ions, and is inhibited by cysteine, cyanide compounds, arsenate various metal chelators, and phosphate ions.

Most conjugates created with AP utilize the form iso-lated from calf intestine.

AP isoenzymes can cleave associated phospho-monoester groups from a wide variety of substrates. The exact biological function of these enzymes is not fully understood, although they definitely function as the opposite of kinase enzymes—removing phosphate groups from phosphorylated proteins and thus affect-ing signal transduction processes within cells. They behave in vivo in their classic phosphohydrolase role at alkaline pH values, but at neutral pH AP isoenzymes can act as phosphotransferases. In this sense, suitable phosphate acceptor molecules can be utilized in solu-tion to increase the reaction rates of AP on selected sub-strates. Typical phosphate acceptor additives include diethanolamine Tris, and 2-amino-2-methyl-1-propanol. The presence of these additives in substrate buffers can dramatically increase the sensitivity of AP-based ELISA determinations, even when the substrate reaction is per-formed in alkaline conditions.

Calf intestinal AP has a molecular weight of about 140,000. The active site of AP contains two zinc ions and a single magnesium ion, both of which are essen-tial for activity (Kim and Wyckoff, 1991). Substrate development with AP should thus be carried out in buffered environments containing small concentrations of these divalent cations to maintain optimal active site conformation. Avoid the presence of metal chelators such as EDTA, since they may extract these ions from the enzyme and inhibit activity. The pH optimum for APs can vary from pH 8 to 10, depending on the type of isoenzyme. Calf intestinal AP peaks in activity at the higher pH values of this range, and substrate reactions are commonly performed in diethanolamine buffer at pH 9.8. The calf intestinal enzyme has the highest cat-alytic rate constant yet discovered for AP isoenzymes, and it is available commercially in high activity for bio-conjugation applications.

Purified preparations of calf intestinal AP main-tained in solution are usually stored in the presence of a stabilizer, which is typically 3-M NaCl. The enzyme may also be lyophilized, but it may experience activity loss with each freeze–thaw cycle. AP is not stable under acidic conditions. Lowering the pH of an AP solution to 4.5 reversibly inhibits the enzyme. It is recommended that all handling, storage, and use of AP be carried out under conditions > pH 7.0 to maintain the highest pos-sible catalytic activity.

AP is often a difficult enzyme to work with when preparing enzyme conjugates. Activity losses may occur upon modification with a crosslinking agent or after coupling to an antibody molecule. Simply fol-lowing established protocols for making antibody–AP conjugates does not always ensure retention of enzyme activity. Sometimes activity losses can be traced to

FIGURE 22.2 Alkaline phosphatase shown as the dimer ribbon structure with two molecules of phosphate bound in its active sites. The molecular model is based on structure 3TG0 in the RCSB Protein Databank by Bobyr et al. (2012).

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particular batches or to certain suppliers of the enzyme. Using a highly purified, high-activity AP prepara-tion helps to maintain good resultant activity in the conjugate.

Ironically, AP is the enzyme of choice for some applications due to its stability. Since it can withstand the moderately high temperatures associated with hybridization assays better than HRP, AP often is the enzyme of choice for labeling oligonucleotide probes (Kaatz et  al., 2012). AP is also capable of maintaining enzymatic activity for extended periods of substrate development. Increased sensitivity can be realized in ELISA procedures by extending the substrate incuba-tion time to hours and sometimes even days, provided that the background interference is low. These proper-ties make AP the second most popular choice for anti-body–enzyme conjugation (behind HRP), being used in almost 20% of all commercial enzyme-linked assays.

Conjugation methods typically employed with AP include glutaraldehyde-mediated crosslinking (Chapter  5, Section 6.2) and the use of the heterobi-functional reagents SMCC (Chapter  6, Section 1.3) or SPDP (N-succinimidyl 3-(2-pyridyldithio)propionate) (Chapter 6, Section 1.1). Heterobifunctional crosslinkers provide the best control over the crosslinking process and typically result in antibody–enzyme conjugates of high activity. Many conjugation protocols incorporate a sodium phosphate buffer system to reversibly block the AP active site during chemical modification. This pre-vents derivatization from occurring in the catalytic site, thus better retaining activity in the resultant conjugate.

1.3. β-Galactosidase

β-Galactosidase (β-Gal; β-d-galactoside galacto-hydrolase; EC 3.2.1.23; also called lactase) catalyzes the hydrolysis of β-d-galactoside in the presence of water to galactose and alcohol. This type of enzyme is found widespread in many microorganisms, plants, and animals. β-Gal can be used to determine lactose in biological fluids and it is employed in food process-ing operations, particularly in immobilized form. The enzyme is often used as a reporter enzyme for moni-toring gene activation and transcription (Beucher et al., 2012; Ju et al., 2012). β-Gal also has good characteristics when conjugated to antibody molecules or streptavidin for use in ELISA systems (Wallenfels and Weil, 1972; Byrne and Johnson, 1975; Kato et al., 1975a,b).

β-Gal has a molecular weight of 540,000 and is com-posed of four identical subunits of MW 135,000, each with an independent active site (Melcher and Messer, 1973) (Figure 22.3). The enzyme has divalent metals as cofactors, with chelated Mg2+ ions required to main-tain active site conformation. The presence of NaCl or dilute solutions (5%) of low-molecular-weight alcohols

(methanol, ethanol, etc.) causes enhanced substrate turnover. β-Gal contains numerous sulfhydryl groups and is glycosylated.

Commercially available β-gal is usually isolated from E. coli and has a pH optimum at 7 to 7.5. By contrast, mammalian β-galactosidases usually have a pH opti-mum within the range of 5.5 to 6; thus, interference from endogenous β-gal during immunohistochemical staining can be avoided.

Due to the relatively high molecular weight of the enzyme, conjugates formed with antibodies and β-gal can be much bulkier than those associated with alkaline phosphatase or horseradish peroxidase. For this reason, antibody conjugates made with β-gal may have more difficulty penetrating tissue structures during immu-nohistochemical or immunocytochemical staining tech-niques than those made with the other enzymes.

Although numerous research articles have been writ-ten describing the preparation and use of antibody con-jugates with β-gal, the enzyme remains a minor player in ELISA procedures. Less than 1% of all commercial ELISA products utilize this enzyme.

β-Gal may be conjugated to antibody molecules using the heterobifunctional reagent SMCC. This cross-linker is reacted first with an antibody through its amine-reactive NHS ester end to form a maleimide-activated derivative. This is in contrast with most anti-body–enzyme conjugation schemes utilizing SMCC, wherein the enzyme is typically modified first and a sulfhydryl-containing antibody is coupled secondarily. However, since β-gal already contains abundant free sulfhydryl residues that can participate in coupling to a maleimide-activated protein, conjugations with this enzyme often are done with the antibody being the first

FIGURE 22.3 β-Galactosidase shown as the four-subunit biologi-cal assembly. The molecular model is based on structure 1BGL in the RCSB Protein Databank by Jacobson et al. (1994).

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component modified. This route avoids having to create sulfhydryls on the antibody molecule, either by reduc-tion or modification with a thiolation reagent. Thus, antibody–β-gal conjugates usually are simpler to make than using other enzymes.

1.4. Glucose Oxidase

Glucose oxidase (β-d-glucose: oxygen 1-oxidore-ductase; EC 1.1.3.4; GO) is a flavoenzyme that cata-lyzes the oxidation of β-d-glucose to d-gluconolactone. The intermediate product of the catalysis is a reduced enzyme–FADH2 complex that, in the presence of oxy-gen, gets oxidized back to enzyme–FAD with release of hydrogen peroxide. The enzyme consists of two iden-tical subunits (MW 80,000 each) bound together by disulfide linkages (O’Malley and Weaver, 1972). GO contains two tightly bound flavin adenine dinucleotide (FAD) cofactors, one per subunit, which are critical to its oxidoreductase activity (Figure 22.4). Each subunit also contains one molecule of chelated iron. The intact protein consists of about 74% amino acids, 16% neutral carbohydrate, and 2% amino sugars (total molecular weight 160,000). GO operates under a relatively broad pH range of 4 to 7, but its pH optimum is 5.5. The com-mercially available preparation of GO is typically iso-lated from Aspergillus niger.

Glucose oxidase is widely used in diagnostic assays for the determination of glucose concentration in physi-ological fluids. Detectability of the oxidation products is done through an enzyme-coupled reaction wherein liberated H2O2 is reacted with peroxidase and a suit-able chromogenic substrate. The development of sub-strate color thus is proportional to the amount of H2O2 released which is in turn related to the amount of glu-cose originally present. The production of hydrogen

peroxide also can be quantified using a luminescence procedure with luminol to produce light in proportion to the glucose concentration (Williams et al., 1976).

GO is often used in solution phase reactions as well as being immobilized on “dip-sticks” and electrodes. Methods for the coupling of enzymes to solid supports can be found in Chapter 15. Although the overall clini-cal usage of GO is widespread, its use as conjugated to antibodies in enzyme-linked assay systems is minor compared to the popularity of other enzymes like HRP and AP. Of the total number of commercial diagnostic assays utilizing antibody–enzyme conjugates, GO is employed in less than 1% of clinical tests. The enzyme remains, however, an important tool in many assays developed for research use (Berron et al., 2011; Holland et al., 2011). One particular advantage to the enzyme is that there is no endogenous GO activity in mammalian tissues, making it an excellent choice for immunohisto-chemical staining procedures.

Antibody conjugates with GO can be made using the crosslinking agents glutaraldehyde (Chapter  5, Section 6.2) or SMCC (Chapter 6, Section 1.3). The het-erobifunctional reagent SMCC provides the best con-trol over the conjugation process and usually results in high-activity preparations. Also consider PEG-based heterobifunctional reagents as described in Chapter 18, Section 1.2, because the hydrophilicity of their cross-bridge provides greater water solubility and less non-specific binding in assay systems.

2. PREPARATION OF ACTIVATED ENZYMES FOR CONJUGATION

Enzymes may be modified to contain reactive groups useful for conjugation with other proteins. This operation may be carried out using homobifunctional (Chapter 5) or heterobifunctional (Chapters 6, 17, and 18) reagents that can covalently couple to some chemical tar-get on the enzyme and result in a terminal reactive group that can crosslink with another molecule. Enzyme acti-vation may also take advantage of the presence of poly-saccharide constituents—oxidizing them with sodium periodate to form reactive aldehydes.

Whatever the method of conjugate creation, the most important considerations are retention of activity in the complex and prevention of extensive oligomer gen-eration, which may cause precipitation. The following methods discuss some of the more common methods for producing enzyme conjugates. The list, however, is by no means exhaustive of every possible procedure used in the literature. Many other crosslinkers and reac-tion strategies as described in this book may be used with success, including the incorporation of polyva-lent scaffolds to attach greater numbers of enzymes to a

FIGURE 22.4 A single subunit of glucose oxidase shown as the ribbon structure with the FADH2 cofactor. The biological assembly typically exists as the dimer. The molecular model is based on struc-ture 3QVP in the RCSB Protein Databank by Kommoju et al. (2011).

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targeting molecule and thus enhance detectability, such as dendrimers (Chapter  8) and polymers (Chapter  18, Section 2).

2.1. Glutaraldehyde-Activated Enzymes

Glutaraldehyde is a homobifunctional crosslinker containing an aldehyde residue at both ends of a 5-carbon chain. Its primary reactivity is toward amine groups, but the reaction may occur by more than one mechanism. As discussed in Chapter 5, Section 6.2, and Chapter 15, Section 2.1, glutaraldehyde is able to cross-link proteins to create stable secondary amine linkages. Glutaraldehyde exists in a number of different forms in aqueous solution, such as hemiacetal and aldol ring structures as well as large α-, β-unsaturated polymers containing carbon–carbon double bonds. The double bonds of these polymers can undergo an addition reac-tion with amines that results in covalent bond forma-tion even without a reductant being present. Fresh glutaraldehyde may contain little polymer formation, except if it is exposed to highly alkaline conditions. However, the older the preparation of glutaraldehyde, the more likely it is that it contains appreciable amounts of polymer. Thus, reactions with this crosslinking agent can result in indistinct conjugation products and may be difficult or impossible to reproduce or scale up as the conjugation need arises.

The high reactivity and indistinct forms of glutaral-dehyde make it difficult to control the size and compo-sition of the final conjugate. Proteins crosslinked with this reagent often form substantial amounts of precipi-tated products due to polymerization. The degree of oligomer formation can be moderated somewhat by using a two-step protocol, but the first protein activated with the molecule can still form large-molecular-weight complexes.

Despite the obvious disadvantages of glutaralde-hyde-mediated conjugation, the crosslinker continues to be used to form enzyme–antibody complexes and to create bioconjugates used in other applications. Many diagnostic tests still utilize antibody–enzyme conju-gates prepared through glutaraldehyde crosslinking procedures.

The one- and two-step procedures for enzyme acti-vation and conjugation using glutaraldehyde can be found in Chapter 20, Section 1.2.

2.2. Periodate Oxidation Techniques

Molecules containing polysaccharide chains may be oxidized to possess reactive aldehyde residues by treatment with sodium periodate. Any adjacent car-bon atoms containing hydroxyl groups will be affected, cleaving the carbon–carbon bond and transforming the

hydroxyls into aldehydes. Glycoproteins may be oxi-dized in this manner to form reactive intermediates useful for crosslinking procedures involving reduc-tive amination (Chapter 4, Section 4). This conjugation technique can direct the coupling process away from polypeptide active regions, thus helping to preserve catalytic activity or binding sites.

Enzymes that contain carbohydrate, such as HRP or GO, may be oxidized with periodate to create reac-tive derivatives that subsequently can be used to label antibodies or other targeting molecules at their amine groups. The aldehyde–HRP intermediate may be stored for extended periods in a frozen or lyophilized state without loss of activity (either enzymatic or coupling potential). Avoid, however, storage in a liquid state, since polymerization may occur—resulting in precipi-tation and loss of activity as Schiff base interactions between proteins builds up in solution.

The protocols for periodate oxidation of HRP and its conjugation with other proteins may be found in Chapter 20, Section 1.3.

2.3. SMCC-Activated Enzymes

The heterobifunctional crosslinker SMCC, or its water soluble analog sulfo-SMCC, can be used to acti-vate enzymes through their amines, leaving terminal maleimide groups on the protein surface (Chapter  6, Section 1.3). The NHS ester end of the crosslinker reacts with ε-lysine or N-terminal amines to form amide bonds. The maleimide end of the reagent is stable enough in aqueous solution to allow purification of the activated enzyme prior to conjugation with a second protein. The maleimide group can react with sulfhydryl groups to create thioether linkages. A maleimide-acti-vated enzyme may be stored in a lyophilized state for extended periods without loss of sulfhydryl-coupling capability.

The use of this type of heterobifunctional reagent allows controlled conjugations to take place, precisely regulating the exact ratio of each protein in the final complex and the size of the resultant conjugate. In addition, the second-stage conjugation through sulfhy-dryl groups provides directed coupling at discrete sites within a protein molecule, thus providing the poten-tial to better avoid active centers or binding regions. For instance, antibody molecules can be coupled to enzymes in their hinge region after mild disulfide reduction to effect the crosslink in an area away from the antigen binding site.

Protocols for the activation of enzyme molecules with SMCC (or sulfo-SMCC) can be found in Chapter  20, Section 1.1. Conjugates formed using this method usu-ally result in high-activity complexes giving excellent sensitivity for use in immunoassays or other appli-

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cations. In addition, refer to Chapter  18, Section 1.2, for the use of similar NHS ester–maleimide crosslinkers that contain a discrete PEG cross-bridge to make the reagent much more hydrophilic. Antibody—enzyme conjugates formed from NHS–PEGn–maleimide reagents perform better in assays than those made from crosslinkers with aliphatic cross-bridges, such as SMCC, because the hydrophobic nature of the aliphatic group often causes nonspecific binding of the conjugate to non-relevant components in a sample. Thus, antibody—enzyme conjugates formed using PEG-based reagents typically have better signal-to-noise ratios in assays, which can be very important when developing diagnos-tic or commercial assays.

2.4. Hydrazide-Activated Enzymes

Hydrazide groups can react with carbonyl groups to form stable hydrazone linkages. Derivatives of pro-teins formed from the reaction of their carboxylate side chains with adipic acid dihydrazide (Chapter  5, Section 8.1) and the water soluble carbodiimide EDC (Chapter  4, Section 1.1) create activated proteins that can covalently bind to aldehyde residues. Hydrazide-modified enzymes prepared in this manner can bind specifically to aldehyde groups formed by periodate oxidation of carbohydrates (Chapter  2, Section 4.4). These reagents can be used in assay systems to detect or measure glycoproteins in cells, tissue sections, or blots (Gershoni et al., 1985).

Other molecules can be used in this type of assay approach, too. Hydrazide-modified (strept)avidin, lec-tins, biocytin, fluorescent probes, and other detectable molecules can be used to detect or image glycoconju-gates in biological samples (Wilchek and Bayer, 1987).

The activation of enzymes using adipic acid dihydra-zide and EDC is identical to the procedure outlined for the modification of (strept)avidin (Chapter 11, Section 5). Simply replace the (strept)avidin component for an equiv-alent mole amount of the enzyme of choice. Alternatively, hydrazide groups may be created on enzymes using the heterobifunctional chemoselective reagents described in Chapter 17, Section 2.

2.5. SPDP-Activated Enzymes

SPDP is a heterobifunctional crosslinker containing an NHS ester on one end and a pyridyl disulfide group on the other end (Chapter 6, Section 1.1). The NHS ester end can be used to modify amine groups on enzymes, forming amide bonds. The result of this procedure is to create sulfhydryl-reactive pyridyl disulfide groups on the surface of each enzyme molecule that are able to complex with thiol-containing proteins and other mol-ecules. SPDP-activated enzymes may be purified and

stored for extended periods without breakdown of the coupling capacity. The reaction with a sulfhydryl group forms a reversible disulfide linkage that can be cleaved with reducing agents.

The two-step nature of SPDP crosslinking provides control over the conjugation process. Complexes of defined composition can be constructed by adjusting the ratio of enzyme to secondary molecule in the reac-tion as well as the amount of SPDP used in the initial activation. The use of SPDP in conjugation applica-tions is extensively cited in the literature, perhaps mak-ing it one of the more popular crosslinkers available. It is commonly used to form immunotoxins, antibody–enzyme conjugates, and enzyme-labeled DNA probes. A standard activation and coupling procedure using SPDP can be found in Chapter 6, Section 1.1.

3. PREPARATION OF BIOTINYLATED ENZYMES

Biotinylated enzymes can be used as detection reagents in (strept)avidin–biotin assay procedures. Particularly, in the bridged avidin–biotin (BRAB) approach or the ABC technique (Chapter 11, Section 2), a biotin-labeled enzyme is used as the signaling agent after the binding to an antigen of a biotinylated anti-body and a (strept)avidin bridging molecule. The biotins on the surface of the enzyme can bind with extraor-dinary affinity to (strept)avidin–antibody complexes, providing near-covalent interaction potential with high specificity.

Adding a biotin label to an enzyme molecule is sim-ple, given the wide variety of options available. A bio-tinylation reagent is chosen that has a reactive group that will couple to functional groups on the enzyme (Chapter 11, Section 6, and Chapter 18, Section 1.3). For instance, NHS–LC-biotin can be used to modify amine groups—a popular choice for many biotinylation pro-cedures involving proteins (Chapter  11, Section 6.2). When free sulfhydryls are present, as in β-gal, a thiol-reactive biotin label may be more appropriate, such as biotin–BMCC (Chapter 11, Section 6.3). However, a bet-ter choice than these popular reagents may be to use hydrophilic biotinylation compounds containing a PEG spacer arm, which results in better solubility of the bio-tinylated enzyme (Chapter 18, Section 1.3). Biotin–PEG reactive compounds are available in different reactivi-ties and spacer lengths to accommodate virtually any application. Enzymes and other proteins modified with these biotinylation reagents retain their water solubil-ity better than when using aliphatic biotin compounds. The hydrophilic PEG spacer prevents aggregation of modified proteins and dramatically reduces nonspecific binding in assays.