bioconjugate techniques || immobilization of ligands on chromatography supports

589Bioconjugate Techniques, Third Edition.DOI: © 2013 Elsevier Inc. All rights reserved.2013http://dx.doi.org/10.1016/B978-0-12-382239-0.00015-7

15

Affinity chromatography has become one of the most powerful techniques available for the isolation of bio-logical molecules. Since the beginnings of this technol-ogy in the late 1960s and early 1970s the application of affinity separations using immobilized affinity ligands has grown to affect nearly every aspect of life science research and has even been used in many other scien-tific disciplines. The basic concept of affinity chroma-tography involves the use of a biospecific or chemically specific interaction between an immobilized ligand and a desired target molecule to selectively bind and interact with the target even in complex solutions con-taining many other components. This specific affinity interaction is able to capture the target while removing contaminants or other molecules in a solution and in a single step enrich or purify the targeted molecule away from all other molecules that cannot bind the ligand (Figure 15.1).

Immobilized affinity ligands have been used for many purposes, including the isolation or purifica-tion of proteins and other biological molecules, in the capture and study of interacting proteins, for removal of contaminants from biological solutions, for enzymatic or chemical catalysis, and for analyti-cal separations involving the assay of a targeted mol-ecule (Calleri et al., 2011; KumaraSwamy et al., 2011; Sheshagirirao et al., 2011; Zeng et al., 2011; Cheung et al., 2012; Vuignier et al., 2012). Chapter 1, Sections 3.3 and 3.4, describe many of these application techniques in the general introduction to bioconjugation. In this chapter, the techniques of immobilization of affinity ligands will be described, including the matrices avail-able for coupling, the activation chemistry used to form reactive groups on these supports, and the cou-pling chemistry commonly used to couple a wide vari-ety of ligand types. Many of the methods are based upon those described in Hermanson et al. (1992), but they are significantly updated here to reflect the latest

developments in chromatography support materials and coupling techniques.

The immobilization of affinity ligands onto insoluble support materials allows the creation of a specific affin-ity matrix that has binding specificity toward a desired target molecule. The insoluble support typically con-sists of a biocompatible material that can be modified to covalently link to the affinity ligand. The word “matrix” often is used to describe any material to which a biospe-cific ligand may be attached covalently. In most cases, the matrix is insoluble in a biological solution contain-ing the target molecule. In some cases, however, the matrix may consist of a soluble polymer that can be modified to contain an affinity ligand.

The most common matrix type used for affinity chromatography consists of porous, beaded materials of generally spherical design that can accommodate biological macromolecules within their porous struc-ture. This type of support material is typically used for chromatographic purposes in the purification of pro-teins and other biological molecules in volumes rang-ing from the bench-top research scale to the bioprocess scale. These materials often are referred to as affinity “supports,” “resins,” “beads,” or “gels.” Affinity chro-matography supports of this type that are used in puri-fication processes often have particle diameters ranging from about 50 to 150 μm and easily settle out of suspen-sion when left standing without mixing. Matrices of this diameter can be used in packed columns or batch operations in scales ranging from microliters or millili-ters for bench chromatography to literally hundreds or thousands of liters in size, which are exploited for many process chromatography applications in biotech and pharmaceutical companies.

In other applications, affinity supports can be prepared using nonporous beads consisting of par-ticles at the low-end micron- or even nanometer-diam-eter range. Affinity supports prepared with such small

C H A P T E R

Immobilization of Ligands on Chromatography Supports

http://dx.doi.org/10.1016/B978-0-12-382239-0.00015-7

BIOCONJUGATE TECHNIQUES

15. ImmobILIzATIon of LIgAndS on CHRomATogRAPHy SuPPoRTS590

particles also can be used in packed columns for chro-matography purposes, such as in HPLC or UPLC sepa-rations, but most often they are used to create insoluble affinity supports for use in batch separations or in ana-lytical techniques. The immobilization of antibodies or other affinity ligands onto microspheres, for example, has been extensively used to develop heterogeneous immunoassays to measure the concentration of pro-teins or other target molecules in biological samples.

Superparamagnetic microspheres can also be coupled with affinity ligands to form a separation system that can capture a target molecule and then be rapidly sepa-rated from solution through the application of a mag-netic field. Such affinity supports are widely used in clinical diagnostic autoanalyzers to measure important analytes in patient samples. See Chapter 14 for a discus-sion on the coupling methods typically used with these very small particles to produce affinity supports.

Sample

The sample is flowedthrough affinity columnin binding buffer Target molecules

bind to affinityligands

Affinity ligands are covalentlyattached to a chromatographysupport and packed in a column

Sample is washed throughthe column to removeunbound components

Purified target moleculeis eluted from the column

Elution buffer is appliedto disrupt the interactionbetween the affinity ligandand the target molecule

1

2

3

4

5

6

FIGURE 15.1 The principal of affinity chromatography in a column format is depicted in this illustration. An immobilized ligand is attached to an insoluble support material which is typically beaded in nature and the affinity resin is packed into a column. The column is equilibrated with binding buffer and a sample containing a target molecule which has affinity for the ligand is passed through the column. The target mole-cule binds to the immobilized ligands and is retained in the column while any not-bound material is washed off. An elution buffer is then added to the column, which disrupts the interaction between the ligand and the target molecule and allows the purified target to be isolated.

5911. SuPPoRT mATERIALS uSEd foR AffInITy CHRomATogRAPHy


1. SUPPORT MATERIALS USED FOR AFFINITY CHROMATOGRAPHY

Affinity supports used for chromatography purposes can be highly diverse and have a wide range of prop-erties. They can consist of natural polymers, synthetic organic supports, inorganic ceramic particles, natu-rally occurring constructs such as diatomaceous earth, membranes or surfaces, or composite constructions consisting of two or more of these materials. The most widely used matrix materials are greater than 10 μm in size and consist of natural polymers such as agarose or synthetic polymeric beads that are formed from hydro-philic monomers. Affinity supports of less than 10 μm in diameter commonly consist of copolymer deriva-tives or silica-based supports. Even extremely small nanoparticles made of metallic spheres or semiconduc-tor alloys also are important substrates for the immo-bilization of affinity ligands; however, the coupling of ligands to nonporous particles of less than about 5 μm in size is described in Chapter 14, while this chapter exclusively focuses on the coupling reactions com-monly used to make affinity chromatography supports on larger porous particles, membranes, and monolithic constructs.

For use with biological molecules, affinity supports must have a base matrix that is both hydrophilic and biocompatible so that the matrix itself does not bind nonspecifically to molecules in a sample. Therefore,

to prepare a highly specific affinity chromatography support, the matrix must have two characteristics that are important to the performance of the final affinity complex: (1) it must be inherently low in nonspecific binding; and (2) it must contain reactive groups or func-tional groups that can be activated to couple the desired affinity ligand. The following sections summarize the major matrix types typically used for affinity chroma-tography and fulfill these criteria for biocompatibility and functionality.

1.1. Natural Product Chromatography Supports

Agarose SupportsOne of the most common support materials is aga-

rose, which is the naturally occurring major polysac-charide component found in agar, extracted from certain red seaweeds (class Rhodophyta, red algae). It consists of alternating residues of d-galactose and 3,6-anhydro-l-galactopyranose (Figure 15.2), with the occasional presence of repeating d-galactose side chains in α-1,4 linkages (initially linked to the C6 hydroxyl of d-galactose of the main chain). Some of the l-galactose residues are not in the anhydro form, thus the poly-mer may contain a quantity of alternating d-galactose and l-galactose residues. The C6 –OH group of the d-galactose units can be methylated in some prepara-tions, as can the C2 –OH group of the l-galactose units on the main chains. The main polymer strands can be

OH

O

OHO

OH

OH OOO

OH

OO

HO

OH

OH OOO

OH

OO

HO

OH

O

OO

O

O

HO

OHO

OHO

HO

OHO

OHO

HO

OHO

OHO

O

HO

OHO

OH

Alpha-1,3 linkage

Beta-1,4 linkage

Alpha-1,3 linkage

Alpha-1,6 linkage

C2 hydroxylspotentiallymethylated

C6 hydroxylspotentiallymethylated

FIGURE 15.2 The repeating disaccharide structure of an agarose polymer is shown, which contains abundant hydroxyl groups that confer excellent hydrophilicity to chromatography supports made of this natural carbohydrate.



further associated together in a left-handed, double- helical construction held together by hydrogen bonds, and these double helices can be further associated with two other double helices in a large wound complex. Two such complexes subsequently can be wrapped around each other into larger supermolecular helices, which are all stabilized by a huge network of hydro-gen bonding between chains. Figure 15.3 illustrates the natural helical arrangement of a single agarose polymer strand. The double and higher order helical polymers arrange themselves together through a complex struc-ture that is extraordinarily gelatinous in nature and when formed into beads. The agarose component takes up only about 2 to 6% of the particles by weight, while the remainder is water which hydrates and swells the aerogel structure. The agarose polymer strands weave throughout each particle to form a highly porous gelati-nous bead that has a translucent appearance under a microscope and a spongy consistency. Since the large hydrodynamic volume of the support forms the bulk of each bead, the pore structure of agarose is huge compared to other chromatography supports. For this reason, agarose can have an exclusion limit for macro-molecules well into the millions of Daltons.

In addition, the carbohydrate nature of an agarose support provides an uncharged, very hydrophilic envi-ronment for optimal use with biomolecules, which naturally has very low nonspecific binding character toward proteins. Commercial beaded agarose sup-ports are typically crosslinked with small bifunctional reagents such as 2,3-dibromopropanol, epichlorohy-drin, or divinyl sulfone to increase the stability of the matrix and prevent melting under elevated tempera-ture or changing structure under denaturing conditions. Non-crosslinked agarose supports also can be used for chromatography, but the chemical, thermal, and mechanical stability of the crosslinked agarose supports is far superior for use in affinity separations.

The aerogel structure of agarose chromatography supports creates a large inner particle environment that accommodates even the very largest macromol-ecules; however, due to its gel-like construction and compressible nature agarose easily can be damaged. The most critical factors that may cause bead damage include mechanical disruption such as grinding par-ticles against surfaces, using too high a flow rate dur-ing chromatography which may result in an excessive pressure drop which crushes the beads, the use of sol-vents that alter the internal structure of the beads by disrupting hydrogen bonding, or drying the particles without the use of an excipient. Agarose actually can be freeze dried and stabilized against particle dam-age through the addition of a sugar molecule such as lactose, which takes up the hydrogen bonding interac-tions upon loss of water. Some reactive agarose sup-ports are stabilized for commercial purposes by freeze drying in the presence of 15% lactose and will rehy-drate rapidly without difficulty or damage to the par-ticles. Other nonreducing sugars may also be used for this purpose, as the multiple hydroxyl groups on the sugars effectively stabilize the agarose polymers and prevent large-scale collapse which could lead to irreversible shrinkage and disruption of the spherical nature of the beads.

Crosslinked agarose can be used with a large number of water-miscible solvents such as dimethyl formamide (DMF), dimethyl sulfoxide (DMSO), tetrahydrofuran (THF), dimethyl acetamide (DMAC), acetone, dioxane, methanol, ethanol, or pyridine, provided the exchange from water into the solvents is done slowly and sequen-tially with increasing concentrations of solvent. The recommended procedure for exchanging agarose into a solvent is to first wash with water, then with 30% of a solvent/water (v/v) solution, followed by a 70% sol-vent/water (v/v) solution, and finally washing the beads with 100% solvent. For complete removal of water from

FIGURE 15.3 A space-filling, twisted helical structure of a single agarose polymer strand is illustrated in this figure. In agarose chromatog-raphy supports multiple double helices are often associated in super-helical structures which include triple helices. These polymer structures are naturally stabilized through numerous hydrogen-bonding sites along the polysaccharide chains. In crosslinked agarose, the structure is fur-ther stabilized by covalent attachments between the chains, which prevent disruption due to melting or denaturation.



within the porous structure of agarose the final wash with solvent should be extensive (at least 10 bed vol-umes of solvent). Many activation methods of agarose are performed in a nonaqueous environment using one of these solvents and it is essential that all water be removed to prevent hydrolysis of the activating agent or hydrolysis of the resulting reactive groups.

The handling of agarose chromatography supports during washing, activation, ligand coupling, and mix-ing procedures should be done carefully to avoid par-ticle damage. Although crosslinked agarose is very robust and can be used with many different reactions, there are a number of caveats that should be considered to avoid gel damage. When filtering agarose to wash with water or solvents, care should be taken to avoid drying out the support during the filtration process. The use of a fritted glass filter funnel using vacuum fil-tration perhaps is the easiest process for filtration and washing, because the particles will not clog the filter and neither will they get lost around a piece of filter paper as they could if a Büchner funnel was used. As the aqueous or solvent solution is filtered through the beads a packed bed of agarose will form on the frit-ted funnel. As the solution filters through the support and reaches the top of the bed, the vacuum should be removed and the filtration stopped to avoid drying out the top of the agarose cake.

In addition, beaded agarose supports should never be used with a stirring bar, because during the mix-ing process the bar can grind the beads against the vessel and cause particle damage. For mixing during activation or coupling procedures, gentle end-over-end mixing or the use of an overhead paddle stirrer should be performed depending on the volume of gel being derivatized. Neither of these methods will cause mechanical damage to agarose particles.

The techniques involved with the immobilization of affinity ligands on agarose are very similar to biocon-jugation reactions used for the attachment of one mole-cule to another. Typically, chromatography supports are handled in bulk using batch methods for washing, mix-ing, activation, or coupling procedures. However, once an affinity support is prepared on agarose by coupling a ligand, any subsequent affinity separations can be car-ried out using batch methods or by column chromatog-raphy while the support is packed in a column.

Agarose chromatography supports can be used with a wide range of activation and coupling reactions. The support mostly contains secondary hydroxyls that originate from the repeating disaccharide composition of the polymer strands along with hydroxyls that may have formed from the crosslinking process if it was done with epoxide-containing reagents. Epoxide cross-linking may also form some primary hydroxyls from unreacted epoxides, which subsequently underwent

ring opening to form diols. In addition, the principal crosslinker used to form crosslinked agarose matri-ces, namely 2,3-dibromopropanol, will add a primary hydroxyl to each site of its reaction within the agarose polymers.

The hydroxyls on agarose may be activated in non-aqueous solution to form reactive electrophilic deriva-tives that are able to react with nucleophiles on affinity ligands to be coupled. Agarose may also be activated in aqueous solution using sodium periodate, which cleaves diols at the associated carbon–carbon bonds to form primary aldehydes. The primary linear chain of agarose only contains diols if some of the 3,6-anhydro-l-galactose are not in the anhydro form and instead exist as l-galactose residues. Since all of the commercial crosslinked agarose chromatography resins can be peri-odate oxidized to a considerable activation level, there must be large amounts of l-galactose present in all aga-rose preparations. Section 2 in this chapter describes many of these activation and coupling reactions along with examples for the immobilization of important affinity ligands.

Agarose chromatography supports are among the most popular beaded matrices for affinity separations. General Electric (formerly Pharmacia and Amersham) manufactures a broad range of basic crosslinked aga-rose beads suitable for separations ranging from bench scale to process chromatography. GE’s agarose sup-ports, sold under the Sepharose name, are the most widely used for biopharmaceutical production, includ-ing the manufacture of recombinant antibodies des-tined for therapeutic use. Other manufacturers, such as Sterogene and Agarose Bead Technologies (ABT), also supply the base gel in one or more forms for use in producing affinity chromatography supports. Many suppliers of affinity media use these base supports to activate and couple affinity ligands. Important com-mercial suppliers of activated agarose supports as well as the associated affinity ligand derivatives include GE, Thermo Fisher, Bio-Rad, Qiagen, and Sigma, among many other companies.

Cellulose SupportsCellulose is a natural polysaccharide polymer con-

taining hundreds or thousands of repeating units of d-glucose linked through β-1,4 glycosidic bonds (Figures 15.4 and 15.5) (Svec, 2002). It is derived from plants, being a primary component of the cell walls, and is commonly used in many consumer products, from biofuels to fiber in foods, paper, and insulation. Cellulose chromatography supports can consist of two main varieties, either an amorphous, fibrous, or crystal-line particulate form with limited porosity or a spheri-cal beaded form having a range of different porosities. Amorphous cellulose has long been used as a rather



crude chromatography support for separations of vari-ous organic molecules and occasionally of biological macromolecules. Its use for affinity chromatography applications has been limited by its poor flow char-acteristics and rather inadequate performance in pro-tein-based separations due to its lack of porosity and capacity.

Cellulose chromatography media are also avail-able in a spherical beaded form with porosities suit-able for affinity separations dealing with proteins and other macromolecules. Natural cellulose supports such as the fibrous variety typically have tightly packed polysaccharide chains, which limit the maximal poros-ity that is achievable. The use of regenerated cellu-lose provides a more hydrated and less tightly packed form of cellulose and thus permits larger pore sizes to be formed when creating beads. Porous cellulose beads are typically made by preparing a solution of a cellulose derivative and dispersing it into another sol-vent system that is immiscible. Rapid stirring of this

mixture forms small droplets that allow the cellulose to re-solidify into spherical beads the size of the droplets. Chisso Corporation (JNC) manufactures a wide range of regenerated cellulose beads under the Cellufine name having two different porosities for use in bio-molecule separations. Some of these supports include a number of immobilized affinity ligands for the sepa-ration of proteins, and they are available in quantities suitable for research chromatography to process scale separations. One matrix in particular, the GCL-2000 gel filtration support, is an underivatized particle having a size exclusion limit of 2 million Daltons, which is emi-nently appropriate for protein separations. This base cellulose matrix may be used in many of the activation and coupling reactions that are described in this chapter for the immobilization of affinity ligands on hydroxylic supports.

Another beaded cellulosic media available for affin-ity chromatography that includes particle sizes ranging from about 30 μm up to 0.5 mm in diameter is produced by Iontosorb in the Czech Republic and distributed under the trade name Perloza. These supports consist of regenerated cellulose with no crosslinks, but nonethe-less they have excellent mechanical stability even with large pore structures, which can accommodate proteins and other macromolecules. Reactive derivatives of the Perloza support are available containing cyanuric chlo-ride or tosyl groups for the immobilization of affinity ligands.

A different type of cellulose matrix that has been used for affinity chromatography separations is avail-able in the form of membranes. Cellulose membranes typically have pore sizes in the low-micron range

OOHO

OH

OH

OO

O

n

Cellulose repeating disaccharide

HOOH

OH

H

FIGURE 15.4 The repeating saccharide structure of cellulose, which contains numerous glucose residues linked through at least two types of glycosidic bonds. Neighboring chains are held together through extensive hydrogen bonding networks to stabilize its macro-molecular structure.

FIGURE 15.5 The three-dimensional structure of a single cellulose polymer is shown, demonstrating the abundant hydroxyl groups that provide hydrophilicity and functional groups for activation and coupling of affinity ligands.



(e.g., 0.45 μm) that easily accommodate proteins and are often used for filtration or clarification of biologi-cal solutions. Cellulosic membranes may be activated in a similar manner to cellulose beads and then cova-lently modified to contain affinity ligands for use in chromatographic separations. Hybrid cellulose mem-branes (containing other polymers) or pure cellulose membranes have both been used for the immobilization of ligands for affinity separations. For instance, a regen-erated cellulose microporous membrane was activated and coupled with protein A to form an affinity support for the purification of antibodies (Boi et al., 2008). The affinity membrane was found to have excellent capac-ity and could capture IgG from polyclonal human anti-body solutions in good yield even at high flow rates. Membranes offer a monolithic alternative to bead-based resins for use in higher flow conditions and they require less maintenance to maintain a column for purification purposes. Although beaded resins still dominate the chromatography industry, membrane-based separations are definitely gaining more interest for their potential advantages in rapid affinity separations.

Inorganic SupportsAnother support material often used for affinity

separations consists of porous inorganic particles of extremely rigid construction and consisting of a variety of possible compositions. Some of the most common matrices in this category include porous glass, silica, alumina, ceramic supports, zeolites, and even the use of naturally occurring siliceous shells of unicellular organ-isms, called diatomaceous earth (or kieselgur; originat-ing from aquatic plants called diatoms).

A commonly used support for affinity chromatogra-phy applications is controlled pore glass (CPG). CPG is typically made from borosilicate glass under high tem-perature resulting in the segregation of the silicate from the borate-rich portions. Subsequent rapid cooling and acidic etching of the particles causes the removal of the borate phase, which in turn results in the formation of craters and pore structures throughout the remaining silicate matrix. CPG supports having pore diameters in the range of ≥1000 Å (≥100 nm) are large enough to allow affinity chromatography for the separation of proteins and other macromolecules. CPG supports are extremely rigid and will hold up to high linear flow rates in packed column beds. However, the glass par-ticles also are breakable, so that care should be taken in handling, mixing, and packing columns, as fines can form easily from particle-on-particle grinding. For this reason, avoid the use of magnetic stir bars or harsh mix-ing conditions. Paddle stirring under moderate mixing rates works well to preserve the integrity of the CPG particles, as does the use of a round-bottom flask in a rotary evaporator under slow rotational speeds.

Another stability issue with glass particles is the potential to dissolve under alkaline conditions. CPG supports are stable at acid or neutral pH, but will dis-solve at pH values equal to or higher than pH 8.0. Buffered conditions containing 50- to 150-mM NaCl should be used for all aqueous conditions with CPG beads, because some degradation can occur even at neu-tral pH in unbuffered water. Conversely, CPG is capable of tolerating a wide range of nonaqueous solvent condi-tions without the shrinking and swelling effects of most polymeric supports. It also withstands high-temperature conditions for use with any derivatization reactions requiring elevated temperatures.

Naked glass particles have considerable potential for nonspecific binding of biomolecules, especially pro-teins and oligonucleotides, due to the preponderance of silicic acid groups (–Si–OH) on the surface. For this rea-son, the base CPG support first needs to be modified to create a layer that is not heavily charged with silicic acid functionalities, which will make it more appropriate for chromatography applications. Typically, this is achieved through silanization of the surface with a functional silane compound that contains uncharged groups and preferentially hydrophilic characteristics. Chapter 13 discusses the modification of particles and surfaces with functional silanes for all kinds of bioconjugation and immobilization applications. CPG beads may be modi-fied with reactive or hydroxylic functionalities to block the silicic acid character and allow subsequent coupling of affinity ligands. Once blocked and functionalized, the degree of nonspecific binding toward proteins and oli-gonucleotides should drop to negligible.

A recommended first step in modifying CPG is to silanize the surface using an epoxy silane, which can be used directly to couple ligands, or alternatively the epoxide groups can be hydrolyzed to form hydrophilic diols (Figure 15.6). The following protocol is based on the methods of Weetal (1969) and Regnier et al. (1976). Similar protocols may be used to coat the particles with other functional silanes containing alternative reac-tive groups or functional groups for immobilization purposes. An important factor in the use of functional silanes to derivatize porous silica or glass particles is that the method must result in only a thin layer of silane coating or the pores will be clogged with silane polymer. In particular, if the reaction is performed in aqueous solution, which is often used for the silane modification of surfaces or nonporous particles, then the resultant high degree of silanol polymerization will quickly reduce porosity. Silylation reactions performed under nonaqueous conditions will limit the degree of modification to only a uniformly thin layer throughout the inner and outer surfaces of the CPG particles, thus preserving the pore structure of the support while effi-ciently coating the raw glass core.



Other silica or inorganic particles containing –OH groups may be modified using the following protocol. For the modification of particles less than about 30 μm, the filtration and washing process should be changed from vacuum filtration to centrifugation or tangential flow filtration to prevent clogging of filters or loss of particles through filter pads that are too coarse to pre-vent the particles from going through the pores.

Protocol1. Add a quantity of CPG particles to a round-bottom,

rotary evaporator flask and add a quantity of 50-mM HCl to cover the matrix and provide enough volume to suspend them during rotation. Place the flask in a

rotary evaporator and slowly initiate rotation while placing the flask under vacuum. The HCl protonates the silicic acid groups and the vacuum removes entrapped air within the pores of the matrix. Mix for 1 h at room temperature, while intermittently breaking and reestablishing the vacuum.

2. Remove the particles from the flask and transfer them to a fritted glass filter funnel or a Buchner funnel containing a glass fiber filter pad. Drain the HCl solution and then wash the particles with 5 volumes of deionized water followed by washing into acetone using sequentially increasing concentrations of acetone in water. Filtration may be carried out by application of a vacuum to a filter

SiO

OO

SiO

OO

SiHO

SiHO

SiOH

SiO

Si

O

SiOH

Si

O

O

SiO

SiO

Si

O

SiOH

Si

O

OH

OH

+

SiO

SiO

Si

O

SiOH

Si

O

NH

R

OH

H2N R

HCl

3-Glycidoxypropyltrimethoxysilane

Controlled poreglass support

containing silanols

Glycidoxypropyl–CPG derivative

Hydrolysis to diol

Coupling of amine-containing ligand

FIGURE 15.6 The reaction of 3-glycidoxypropyl trimethoxysilane with CPG and hydrolysis of the epoxide groups to yield diols or coupling of an amine-containing ligand to the epoxy groups to yield a secondary amine linkage.



flask to increase the flow through the filter pad. Wash with at least 10 volumes of pure acetone to remove the last traces of water. Allow the particles to dry on the filter while applying a vacuum to aid in the removal of acetone by filtration and evaporation. Gently break up the filter cake using a plastic spatula to better disperse the particles and facilitate drying.

3. Transfer the particles to the round-bottom rotary evaporator flask and cover them in toluene. Again rotate the flask for an hour while pulling a vacuum to remove entrapped air. Next, heat the slurry in the rotary evaporator unit to 40 to 60°C while continuing the application of vacuum to begin distillation of the toluene. Continue the distillation process until water stops azeotropically distilling along with the toluene. If the last traces of water are not completely removed, the silylation process will result in hydrolysis of the trimethoxy groups and rapid polymerization of the functional silane, which will clog the pores of the CPG support. Cool the flask and remove excess toluene from the support by filtration.

4. Prepare a solution containing 10% (v/v) 3-glycidoxypropyl trimethoxysilane (Acros Organics) and a total volume equal to the volume of particles to be treated. Transfer the support back into the rotary evaporator flask and add the silane solution to the particles to resuspend them. React overnight at room temperature with rotation at slow speed.

5. Transfer the particles to the filter funnel and remove excess silane solution. Wash extensively with dry acetone to remove remaining traces of silane and toluene. Finally, filter off the acetone and dry the particles under vacuum. The coated, epoxy functional particles may be used directly for coupling an affinity ligand or may be treated as in step 6 to hydrolyze the epoxide group to a hydrophilic diol. Protocols for the coupling of amine-, thiol-, or hydroxyl-containing ligands to epoxide-reactive groups may be found elsewhere in this chapter.

6. Transfer the modified CPG support into a clean rotary evaporator flask and add a quantity of 50-mM HCl to cover the particles. Place under vacuum to remove entrapped air with slow rotation. Heat the flask to 50 to 60°C for 1 to 2 h to hydrolyze the epoxide groups into diols. The hydroxyl groups may be used in a number of different activation reactions to couple affinity ligands. For instance, the adjacent hydroxyl groups may be oxidized with sodium periodate to yield a reactive aldehyde group for coupling to amine-containing molecules.

Glass supports treated in this manner to yield a diol coating on the surface will have low nonspecific bind-ing to proteins or other biomolecules. The silylation process coats the inner and outer surfaces of the

particles and prevents biomolecule interactions with the silicic acid functionalities. However, it is known that silica- and glass-based chromatography supports give higher potential nonspecific binding character toward protein-containing samples than agarose supports, pre-sumably due to some exposure of the silica backbone (Ghose et al., 2007). Alternative silane treatments may also be carried out with CPG supports to create other functionalities for further derivatization. In particu-lar, an aminopropyl silane may be used to add amine functionalities to a glass surface, which may be used for direct coupling of carboxylate-containing ligands or to build hydrophilic spacer arms prior to coupling an affinity molecule. In addition, other support materials such as silica and alumina may be treated in a similar manner to create functional matrices suitable for affin-ity chromatography. See Chapter 13 for further informa-tion regarding functional silane compounds and their reactions with substrates.

Commercial affinity supports prepared on glass par-ticles include a high-capacity protein A or protein G matrix, which has a high density of immunoglobulin-binding proteins immobilized on its surface (Prosep-A or Prosep-G from Millipore or Trisopor-Protein A from BioCat). These supports can tolerate high flow rates during chromatographic purification of antibod-ies, because the base particle is extremely robust and incompressible. Binding capacities of over 45 mg/ml for antibodies are not unusual on this type of affinity sup-port material.

Composite Gel–Ceramic SupportsNovel chromatographic support materials have also

been created from the use of porous, inorganic particles that have been coated with a gelatinous natural poly-mer. The core ceramic particle provides an incompress-ible substrate that can take the rigors of large columns and rapid flow rates, while the gel coating creates a hydrophilic surface that is low in nonspecific binding character. This type of support was originally developed by BioSepra and commercialized under the trade name HyperD. BioSepra subsequently underwent a series of acquisitions and sales until the company was ultimately sold to Pall Corporation in 2004. U.S. patents 5,234,991 (1987) and 5,268,097 (1993) describe the preparation and use of these “gel-in-a-shell” supports, which consist of a base inorganic mineral oxide bead containing a hydro-gel-filled pore structure (Figure 15.7). The type of core particle used for the HyperD supports is a porous silica bead called Spherosil, which was originally manufac-tured by Rhone-Poulenc but is now produced by Pall Corporation.

The hydrogel that is used to fill and coat the inter-nal surfaces should ideally be a long linear polymer with a molecular weight of at least 104 Daltons. It also



should be hydrophilic to produce a biocompatible surface and be partially cationic in nature in order to stick to the core inorganic support material, which has a negative charge character. Most often the polysac-charide derivative is aminated to contain tertiary or quaternary amine derivatives along the length of the polymer. Examples of polymers that can be used in this fashion include DEAE–dextran and QAE–dextran derivatives, which contain multiple positively charged amines. It is believed that the positively charged poly-mer interacts with the negatively charged mineral oxide bead surface and forms multi-point, strong noncova-lent interactions that are essentially irreversible. For instance, in the modification of CPG or silica supports, the negatively charged silicic acid groups likely inter-act with the cationic groups of the polymer, creating a large number of charge interactions along each poly-mer molecule. The polymer-coated surface also can be treated with a crosslinker such as a bis-epoxide or epi-chlorohydrin to further stabilize the hydrogel and pro-vide a composite gel–ceramic particle, which is stable to strong acid or base without disrupting the polymer coating.

The use of aminated dextran, starch, or agarose as the gel layer for coating the particles can result in a poly-hydroxylic support material that can be activated to contain reactive groups capable of coupling affin-ity ligands. Methods to produce this type of composite gel–ceramic particle can be used successfully on such mineral oxide supports as glass, silica, alumina, tita-nium, and magnesium. The final “gel-in-a-shell” con-struction combines the benefits of the rigid ceramic core with the hydrophilicity and immobilization capacity of the biocompatible polymer (Xia et al., 2011). The inor-ganic phase typically consists of particles with huge

pore structures that allow for fast mobile-phase access that eliminates the diffusion limitations often seen with standard porous beaded supports. The large pores allow rapid sample distribution throughout the interior structures of the beads, thus overcoming the drop-off in capacity usually observed as flow rates through a col-umn increase.

Activation and coupling methods for the immobili-zation of affinity ligands on these composite supports can involve any of the methods described in this chap-ter for the activation of hydroxylic functional groups. This includes the use of aqueous periodate oxidation to form aldehyde groups, which can then be used to cou-ple amine-containing ligands such as proteins through the use of a reductive amination reaction process. This activation and coupling reaction process is described elsewhere in this chapter in the section entitled Amine-Reactive Immobilization Methods.

1.2. Synthetic Polymeric Chromatography Supports

Perhaps the most diverse category of affinity chro-matography supports is represented by resins that con-sist of various types of synthetic polymers. Synthetic supports are produced from the controlled polymeriza-tion of various functional monomers and crosslinking monomers in a stirred solution that produces beaded particles having a porous structure. In some cases, poly-meric supports may consist of a combination of natural polymers and synthetic polymers. For instance, an aga-rose–polyacrylamide composite support may be made that combines the hydrophilic functionality of a poly-saccharide with the tight pore structure of a polyacryl-amide particle, thus creating a matrix that can be easily derivatized to contain affinity ligands.

Synthetic polymeric supports can be designed to have a variety of different physical characteristics that are appropriate for chromatographic separations (Gokmen and Du Prez, 2012). As compared to natu-ral supports, synthetic matrices may have superior physical and chemical stability, while still being able to withstand the high flow rates typically seen in pro-cess chromatographic separations. Polymeric supports also can withstand enzymatic degradation or microbial contamination better than some naturally occurring polymers, which may be ideal substrates for bacterial growth. Porous polymeric chromatography supports designed by using reactive or functional monomers can easily be coupled directly with affinity ligands or activated to contain a reactive group for immobiliza-tion. Many commercially available synthetic polymeric supports contain primary or secondary hydroxyls and are extremely hydrophilic and low in nonspecific bind-ing. In addition, hydroxylic-containing supports can

Ceramiccore particlePolymer-filled

pores

FIGURE 15.7 An illustration of a gel–ceramic composite sup-port material. The ceramic core forms the structural spheroid shape of the support, and it has also very large pore structures to accom-modate macromolecules. These pores are filled with polymeric gels, such as agarose, to create a hydrophilic porous resin, which combines the advantages of the polymer with the robust and rigid nature of the ceramic core to give a superior chromatographic support useful for process scale affinity chromatography.



be activated by a number of different immobilization chemistries and subsequently coupled with a wide vari-ety of affinity molecules.

Polyacrylamide SupportsPolyacrylamide chromatography supports are typi-

cally made by the copolymerization of two monomers: acrylamide and N,N’-methylene bisacrylamide, first described by Hjerten and Mosbach (1962) (Figure 15.8). The support can have a variety of pore sizes depend-ing on the degree of crosslinking monomers added to the polymerization reaction. The greater the amount of bisacrylamide present in polyacrylamide, the smaller the pore structure in the resultant beads. Bio-Rad offers a broad range of polyacrylamide resins that differ in the pore size and exclusion limit of the particles. These sup-ports are classic gel filtration (size exclusion chromatog-raphy, or SEC) matrices that can separate proteins based on their molecular weight. However, they may also be used as supports to couple affinity ligands for purifi-cation of selected target molecules. For working with proteins, a size exclusion limit for the polyacrylamide beads should be chosen that will allow the protein mol-ecules to enter the pores and interact with immobilized affinity molecules.

Polyacrylamide resins are typically supplied dry and must be hydrated in aqueous solution with gentle agita-tion prior to use. Weigh out an appropriate amount of dry beads and slowly add them with mixing to a quan-tity of aqueous buffer equal to about twice the antici-pated volume of hydrated gel.

Polyacrylamide supports are characteristically soft in nature and gelatinous in appearance. They also dis-play low nonspecific binding character toward biomol-ecules, rather good pH and buffer stability, and because they are totally synthetic, the supports are not able to support microbial growth. In addition, since polyacryl-amide does not contain carbohydrate components, the support is ideal for use in the affinity purification of carbohydrate-binding proteins. For this reason, affin-ity supports prepared on polyacrylamide containing immobilized sugars, disaccharides, or small polysac-charides often have been used for the specific purifica-tion of lectins.

A major deficiency of polyacrylamide-type resins is their inability to support acceptable flow rates to per-mit larger-scale separations to take place in a reasonable time frame. Due to the compressibility of polyacryl-amide, maximal linear flow rates are often limited to no more than 10 to 15 cm/h, which is far less than 10% of the flow rate that can be expected from agarose supports or other polymeric matrices. In addition, polyacrylamide beads have a tendency to shrink and swell with different solvents, buffers, and salt concentrations. Severe reduc-tion in particle size can occur in organic solvents and

this characteristic limits the range of chemical reactions that can be done to immobilize affinity ligands on them. Shrinkage of the particles also causes a collapse of the internal pore structures, which in this case may restrict coupling of ligands only to the outer surfaces.

Chromatography particles made of polyacrylamide do not naturally contain reactive groups or functional groups suitable for the immediate attachment of affin-ity ligands. Derivatives of polyacrylamide are typically made through partial hydrolysis of the amide bond linkages within the gel or through a process of trans-amidination carried out at elevated temperatures using amine-containing spacer arms or ligands. Using such strategies a number of methods have been developed to produce the requisite functional or reactive groups needed for immobilization of a wide range of affinity ligands.

Modification of polyacrylamide through transam-idination involves the addition of a high concentra-tion of an amine-containing ligand and heating while gently mixing. For instance, a diamine-containing spacer molecule may be reacted in large excess with the polyacrylamide support while mixing at 90°C and the amide bonds within the matrix will be broken and reformed with a linkage to the amine group on the

NH2

O

NH

O

NH

O

+

NH2

O

NH2

O

NH

O

NH2

OH2N O

NH2O

HN O

NH2O

H2N O

O

H2N O

H2N O

NH2

O NH2

O NH2

O NH2

O

HN

H2N O

O

H2N O

H2N O

NH2

O NH2

O NH2

O HN

O

H2N

Acrylamide Methylenebisacrylamide

PersulfateTEMED

Crosslinkedpolyacrylamide matrix

FIGURE 15.8 Polyacrylamide supports are made from the copo-lymerization of acrylamide and the crosslinking monomer N,N’-methylene bisacrylamide. The ratio of crosslinking monomer to acrylamide monomer controls the degree of porosity in the final sup-port material.



spacer (Figure 15.9). A high density of such modifi-cations may be obtained using a variety of amine- or diamine-containing molecules. In addition, hydrazide groups may be formed on the polyacrylamide support by reacting the gel at 50°C with an excess of hydrazine. (Note: Hydrazine is extremely toxic and explosive—use a fume hood and use proper personal protective equip-ment to handle safely.)

The following protocol describes the modification of polyacrylamide beads with ethylenediamine (EDA) using thermal transamidination, as disclosed by Inman and Dintzis (1969). A similar procedure may be used to create hydrazide groups using hydrazine substitution. All operations using EDA or hydrazine should be done in a fume hood using appropriate protective equipment.

PROTOCOL

1. Set up a three-necked, round-bottom flask in a fume hood with a heating mantle and paddle stirring rod and overhead stirring motor. Add 500 ml of EDA to the flask and stir the solution while heating to 90°C. Monitor the temperature using a thermometer inserted into one of the three necks in the flask.

2. Stir the EDA solution rapidly while slowly adding 25 g of dry polyacrylamide beads. The beads should be added in small amounts to avoid clumping of the particles. The beads will swell in the hot EDA solution as the reaction occurs.

3. Stir the solution for 5 h at 90°C.4. Remove the heating mantle and allow the gel slurry

to cool to room temperature with continued stirring. After cooling, add an equal volume of crushed ice made from deionized water to chill the slurry as it comes into contact with aqueous solution, because the EDA solution is highly basic and its dissolution into water is exothermic.

5. Transfer the gel slurry with the crushed ice to a fritted glass filter funnel or Büchner funnel containing a glass fiber filter pad and wash the matrix with 0.2-M NaCl in 1-mM HCl. Add additional crushed ice to prevent severe heating of the gel during the initial wash steps. Wash the gel until the excess EDA has been removed and the gel has decreased in pH to neutrality or acidic conditions. After the unreacted EDA has been washed out of the support, wash thoroughly with 0.2-M NaCl for at least 5 bed volumes. The TNBS test for amines may be used to check for the presence of EDA in the washings and to check a small volume of derivatized gel for successful coupling (Chapter 2, Section 4.3). An orange color is indicative of the presence of amines.

Trisacryl SupportsAnother polymeric support made for chromato-

graphic operations is Trisacryl, a matrix that was origi-nally produced by IBF Biotechnics but is now available from Pall Corporation as a result of several acquisitions. The polymeric support gets its name from the fact that a key monomer used in the copolymerization pro-cess is made from the amide derivative of acrylic acid with Tris buffer (tris(hydroxymethyl)aminomethane), the actual chemical name for which is N-acryloyl-2-amino-2-hydroxymethyl-1,3-propanediol. This acryl-amide derivative is reacted along with the crosslinking dimer N,N’-diallyltartradiamide or another crosslink-ing dimer, N,N’-methylene bisacrylamide, to give the final beaded, porous polymer known as Trisacryl (Figure 15.10). The Tris component of the polymeric mixture provides strong hydrophilicity due to the three

NH2

O

NH2

O

NH

O

NH2

OH2N O

NH2O

H2N O

NH2O

H2N O

O

H2N O

H2N O

NH2

O NH2

O NH2

O NH2

O

HN

90° C

NH2

O

NH2

O

NH

O

NH2

OHN O

NH2O

HN O

NH2O

H2N O

O

H2N

H2N

O

H2N O

HN

O NH2

O HN

O NH2

O

HN

NH2

NH2

NH2

NH2

NH2

Polyacrylamidematrix

Ethylenediamine

Polyacrylamidecontaining

primary amines

FIGURE 15.9 Polyacrylamide supports may be modified with ethylenediamine using a transamidination reaction at high tempera-ture to produce amine groups on the support for further reactions.



hydroxyl groups present on each monomer unit, while the crosslinking monomer is added in an optimized ratio to control the final porosity and mechanical strength of the particles (Batista-Viera et al., 2011). The main Trisacryl polymers are highly dense structures three-dimension-ally with an outer presentation of hydroxyl groups completely surrounding each linear chain and periodic crosslinking monomers bridging the chains to create a stable matrix (Figure 15.11).

The primary hydroxyls within Trisacryl create abun-dant functional groups for activation and coupling of affinity ligands. They also help to prevent nonspecific binding to biological molecules, thus making affin-ity supports made on Trisacryl have the potential to be highly specific without secondary “matrix effects.” Limiting the degree of crosslinking within the matrix can produce a support with large pores having an exclusion limit exceeding 10 million Daltons, which makes it imminently fitting for protein separations. This type of beaded support, called Trisacryl GF-2000,

is available in two particle size ranges that can be cho-sen to reflect the needs of the affinity chromatography application. The smaller particle size range, 40 to 80 μm, can be used for analytical to medium-scale separations wherein tighter peak shapes are desired from packed columns used along with bench-scale liquid chroma-tography systems, for instance FPLC-type instruments. Alternatively, a larger particle size range is available, 80 to 160 μm, that can be used for process scale separa-tions where faster flow rates are necessary for greater efficiencies in the isolation of biomolecules. Both sup-ports have identical porosities, so the ultimate choice of particle size should be made based upon the intended application needs.

Unlike plain polyacrylamide resins described pre-viously, Trisacryl supports have much greater rigid-ity and are not as gelatinous in nature. For this reason, columns packed with Trisacryl have better stability to changes in mechanical pressure and flow rates used during chromatography; therefore affinity resins made using this support can be used without difficulty to pro-duce large quantities of proteins or other biomolecules in process separation procedures. The chemical sta-bility of Trisacryl also is excellent, as it can be used in pH environments from pH 1 to 11 without breakdown due to hydrolysis. At highly basic pH values, however, the support material may hydrolyze by cleavage of the polymer amide bonds; therefore, only brief exposure to NaOH (0.2 N) for sanitization and regeneration should be done to prevent matrix damage.

The abundant hydroxyl groups within Trisacryl can be used for activation and immobilization of affinity ligands. The activation reactions typically form inter-mediate electrophilic reactive groups that can be used to couple with affinity ligands containing nucleophilic groups, such as amines, thiols, and other hydroxyls. Examples of activation reagents that can be used with Trisacryl include cyanogen bromide (CNBr), carbonyl

NH

OOH

OH

OH

NH

HN

O

O

OH

OH

N-Acryloyl-2-amino-2-hydroxymethyl-1,3-

propane diolN,N '-Diallyltartradiamide

+

HN O

OHHO

HO

HN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HO

HN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HO NH

HN

O

O

HO

OH

HN O

OHHO

HO

HN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HO

HN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HOHN O

OHHO

HO

CrosslinkedTrisacryl Polymers

FIGURE 15.10 Trisacryl supports are made by the copolymeriza-tion of a vinyl monomer containing a tris-hydroxymethyl group and the crosslinking monomer N,N’-diallyltartradiamide. The support contains a high density of hydrophilic hydroxyl groups, which can be used for activation and immobilization reactions to couple affinity ligands.

FIGURE 15.11 The three-dimensional, space-filling molecular model of a portion of the Trisacryl polymeric structure showing a sin-gle crosslink.



diimidazole (CDI), tresyl chloride, tosyl chloride, divi-nyl sulfone, and bis-epoxides such as 1,4-butanediol diglycidyl ether (see corresponding sections later in this chapter). The only caveat to keep in mind with regard to the activation of Trisacryl is that sodium periodate treatment to form aldehydes, which is a common route to form reactive groups on polysaccharide-containing supports, cannot be used successfully. This is due to the fact that the hydroxyls on Trisacryl come from the

Tris groups along the polymer strands and are not on adjacent carbon atoms, so there are no periodate-oxidiz-able groups present within the matrix. One alternative strategy to create aldehyde groups that has been found to be effective is to first react the matrix with glycidol (a monofunctional epoxide–alcohol; Figure 15.12), and then periodate-oxidize the resultant gel, which now contains diols, to form reactive aldehydes (see corre-sponding section, this chapter).

Sephacryl SupportsSephacryl supports were originally developed by

Pharmacia and are now sold by GE Healthcare Life Sciences as a result of a series of company acquisitions and mergers. This support is actually a composite of a naturally occurring polysaccharide derivative and a synthetic polymer. The porous, beaded support is made from the copolymerization of allyl dextran and N,N’-methylene bisacrylamide wherein the dextran carbohy-drate is crosslinked by the bisacrylamide, which forms a strong hydrophilic matrix consisting of a multitude of glucose residues (Figure 15.13). The relative porosity of the matrix can be controlled by the amount of the com-ponents, especially the crosslinking monomer, added to the polymerization reaction. Sephacryl is available in a range of different molecular weight exclusion limits, and the largest of these, Sephacryl S-300 or S-400 with exclusion limits of 1500 and 8000 kDa, respectively, are appropriate for use as affinity supports as they can accommodate interacting proteins within their pore structure without difficulty.

Some Sephacryl supports have particle sizes that are rather small for use in gravity columns or for process-scale separations, because the matrix was originally designed to be a gel filtration or size exclusion type of support, which provides high resolution on FPLC- or HPLC-based applications. However, the Sephacryl HR series of supports has a particle size distribution that is larger than the Sephacryl SF series and this allows it to be used for moderate-scale affinity separations (25- to 75-μm diameter particles).

Sephacryl supports are supplied pre-swollen as an aqueous suspension containing a preservative. The gel is relatively stable to mechanical stresses during han-dling, so it can be mixed and washed using standard fil-ter funnels to exchange the matrix into the appropriate solvent or buffer for activation and coupling. Stirring should be performed using a paddle stirrer or by rota-tion, however, and not using a stir bar to avoid grind-ing and breakdown of the support. Also avoid drying the support to prevent collapse of the particle, which may be irreversible without the presence of an excipient such as lactose in high concentration.

Immobilization reactions performed on dextran-based gels like Sephacryl can take advantage of the

NH

O

HO

OH OH

Trisacryl polymercontaining hydroxyl groups

+ OHO

Glycidol

NH

O

OH

O

OHOH

O

OHO

OH

O

OHOHO O

OHOH

O

OHOH

O Branched glycidolpolymer modifications

NH

O

OH

O

H O

O

OHO

OH

O

OHOHO O

H

OO

H O

O Oxidation of terminaldiols to aldehyde residues

+ 3H2CO

Sodiumperiodate

FIGURE 15.12 Glycidol modification of Trisacryl supports fol-lowed by periodate oxidation to form aldehydes useful for coupling amine-containing affinity ligands. The glycidol polymers formed within the porous structure of the support can be controlled as to their density and size by modulating the mole excess of glycidol added to the reaction.



abundant quantity of hydroxyl groups on the glucose residues within the matrix. Since the bisacrylamide crosslinks provide increased physical stability, the sup-port will tolerate washing into environments including 100% organic solvents, which are often used for acti-vation reactions prior to coupling an affinity ligand. In this regard, activation reactions using CDI, tresyl chloride, and tosyl chloride can be carried out to cre-ate intermediate reactive groups that then can be used in organic solvent or transferred back into aqueous buf-fers for coupling to amine-containing ligands. In addi-tion, aqueous-phase activation methods also can be used with Sephacryl, including periodate oxidation of the diols on glucose to form aldehydes, activation with divinyl sulfone, or activation with bis-epoxide-based

compounds such as 1,4-butanediol diglycidyl ether. See the corresponding sections in this chapter for the exper-imental protocols related to activation and coupling using these methods.

Ultrogel AcA SupportsOriginally, Ultrogel supports were produced by

IBF but as a result of acquisitions they are now sold by Pall Corporation. The supports consist of agarose that is copolymerized with acrylamide monomers to produce a particle that has the biocompatible advan-tage of agarose and the size exclusion and controlled porosity capability of polyacrylamide-based supports (Figure 15.14). The result is a range of size exclusion matrices that are superior to standard polyacrylamide

O

HOHOOH

O

O

HOHOOH

O

O

HOHOOH

O

O

HOHOO

O

OAllyl dextran

polymer

+ NH

O

NH

O

Methylene bisacrylamide

O

HOHO

OHO

O

HOHO

OHO

O

HOHO

OHO

O

HOHO

OOO

O

HOHO

OHO

O

HOHO

OHO

O

HOHO

OHO

O

HOHO

OOO

NH

NHO

O

Crosslinked dextran

HN

O

HN

O

NH

HN

O

O

O

HO

HO

HO

O

O

HO

HO

O

O

O

FIGURE 15.13 The chemical nature of Sephacryl supports, which are formed from the copolymerization of N,N’-methylene bisacrylamide and allyl dextran to create a crosslinked dextran matrix having good physical stability.



H2N O

O

H2N O

H2N O

NH2

O NH2

O NH2

O NH2

O

H2N

H2N O

O

H2N O

H2N O

NH2

O NH2

O NH2

O NH2

O

H2N

H2N O

O

H2N O

NH2

O NH2

O NH2

OH2N

NH2

O

NH2

O

NH2

OH2N O

NH2O

H2N O

NH2O

NH2

O

NH2

O

NH2

O

NH2

OH2N O

NH2O

H2N O

NH2O

H2N O

O

H2N O

NH2

O NH2

O NH2

OH2N

H2N O

NH2

O

H2N O

O

H2N O

H2N O

NH2

O NH2

O NH2

O NH2

O

H2N

H2N O

O

H2N O

H2N O

NH2

O NH2

O NH2

O NH2

O

H2N

H2N O

O

H2N O

NH2

O NH2

O NH2

OH2N

OH

OH

OHO

HOOH

HO

O

O

OOH

O

OHO

OH

HO

O

O

OOH

O

OHO

OH

HO

O

O

OOH

O

OHO

OH

HO

OH

O

OOH

O

OHO

HOOH

HO

O

O

O

O

OHO

OH

HO

O

O

OOH

O

OHO

OH

HO

O

O

OOH

O

OHO

OH

HO

OH

O

O

O

OHO

HOOH

HO

O

O

OOH

O

OHO

OH

HO

O

O

OOH

O

OHO

OH

HO

O

O

OOH

O

OHO

OH

HO

OH

O

OOH

O

Agarose

Polyacrylamide

FIGURE 15.14 Ultrogel supports contain a combination of agarose polymers and polyacrylamide to create a complex network within the particles.



supports, but have smaller pore structures than sup-ports made of agarose alone. In the preparation of the beads, the percentage of polyacrylamide in the sup-port governs the final porosity, with a higher percent-age of agarose providing a higher exclusion limit of the final support. A particular Ultrogel support contain-ing 2% agarose and 2% polyacrylamide provides an exclusion limit of about 3000 kDa, which is sufficient for most affinity chromatography applications involv-ing proteins. Unfortunately, Ultrogel supports do not utilize crosslinked agarose in the preparation and thus they may have some solvent sensitivity in addition to the potential to be damaged by high concentrations of denaturants or extremes of pH.

Ultrogel is also available as a paramagnetic particle in which iron oxide (Fe3O4) has been incorporated into the core of the polymer. The resultant bead can be used in magnetic separations wherein the particles can be removed from solution by the application of a mag-netic field. The porosity of this particle is not as great as that of the chromatographic support in that it only has an exclusion limit of 200 kDa, which will likely reduce the overall capacity of the matrix for binding proteins. However, it will likely have superior binding capacity compared to the use of nonporous magnetic particles, which are often used in small-scale affinity separations.

Ultrogel supports should not be used with activa-tion methods that require nonaqueous conditions, as solvents will damage the support. However, the matrix may be activated in aqueous conditions using CNBr, divinyl sulfone, or epoxide activation methods with suc-cess. It is anticipated that activation with bis-epoxides will create internal crosslinks and help to stabilize the support as compared to the base support. See the appro-priate sections in this chapter that describe these activa-tion protocols.

UltraLink Azlactone SupportsThe UltraLink resins that are available from Thermo

Fisher Scientific (Pierce) were originally developed at 3M in the mid-1990s and commercialized under the Emphaze name. This support material consists of the copolymerization of vinyldimethyl azlactone (oxazo-lone) with the crosslinking monomer, N,N’-methylene bisacrylamide (Figure 15.15). The azlactone monomer creates a particle containing reactive groups that is immediately useful in the coupling of amine-contain-ing ligands, such as proteins. Therefore, the base sup-port is unusual in that it comes already activated for the immobilization of affinity ligands, which is a feature designed into the matrix from the start. The amount of azlactone-reactive groups can be controlled simply by altering the ratio of the reactive monomer to the cross-linking monomer. Beads have been synthesized that contain from 100 μmoles/ml gel to 300 μmoles/ml of

azlactone functionality. The typical azlactone reactivity in UltraLink resin is sufficient for the immobilization of over 30 mg/ml of protein A or over 20 mg/ml of human IgG, which is in excess of the levels usually required for protein-based affinity ligands.

The UltraLink support in its active form comes as a dry powder that, upon addition to an aqueous buffer, swells to become fully hydrated beads with a particle size of about 50 to 80 μm and a spherical appearance under a microscope. The beads typically hydrate in aqueous solution to yield a support having a total swollen volume of about 10 ml/g of particles. The fully swollen particles have pore structures of at least 500 Å with a total surface area of about 250 m2/g. The porous internal structure of the beads is large enough to accommodate interacting proteins, including antibod-ies, thus making the support very appropriate for affin-ity chromatography applications dealing with proteins. Once the particle is hydrated, the azlactone-reactive groups are susceptible to hydrolysis and so they should be added directly to a ligand solution for coupling to avoid losing reactivity. Although the azlactone groups will hydrolyze with ring opening to terminal carboxyl-ates (Figure 15.16), the rate of hydrolysis is relatively slow compared to other electrophilic groups com-monly used for immobilization. The rate of coupling to an amine-containing ligand takes place with maximal yield usually within 1 to 2 h at room temperature.

Azlactone groups are reactive toward nucleophiles (e.g., –NH2, –SH, –OH), but with the greatest cou-pling potential being with amines at basic pH val-ues. The reaction proceeds with nucleophilic attack on the azlactone carbonyl group causing ring opening and rearrangement to an amide bond linkage with the amine-containing ligand and forming a short spacer arm to the matrix (Figure 15.17). Small amine-con-taining ligands may be coupled in aqueous or organic solution, provided that the organic solvent chosen allows the beads to fully swell upon addition to the ligand solution, so that the internal pore structures may be accessed during the immobilization reaction (see Table 15.1).

The UltraLink polymeric support has excellent mechanical properties for rapid chromatographic sepa-rations, being able to tolerate high linear flow rates while still maintaining high binding capacities. The support will tolerate flow rates several times higher than highly crosslinked agarose supports, thus mak-ing it useable in large columns under rapid flow with-out collapse of the particles. The particle structure permits high-pressure chromatography, with typical maximal pressure drops across the column exceeding 1000 psi before bead damage is observed. In practice, such extreme pressure conditions are never required, because very high flow rates can be realized under



pressures of far less than 100 psi. In addition, stud-ies performed with derivatized UltraLink supports have demonstrated the excellent chemical stability of the base copolymer construction as well as the stabil-ity of the covalent bond formed with amine-containing ligands. The polymeric matrix is stable to degradation in extremes of pH (1–14) and heat, as well as being

resistant to harm in highly denaturing conditions or high salt or buffer conditions.

For use of UltraLink azlactone-activated supports in the coupling of affinity ligands, the dry matrix should be accurately weighed to result in a known volume of hydrated gel in the coupling buffer. Approximately 0.1 g of beads will be hydrated to a volume of 1 ml of gel,

NH

O

NH

O

N,N '-Methylene bisacrylamide

N

O O

Vinyldimethyl azlactone

+

N

OO

N

OO

N

OO

N

OO

N

OO

N

OO

N

O

ON

O

ON

O

ON

O

ON

O

ON

O

O

NH

O

NHO

N

O

O

N

O

O

N

O

O

N

O

O

N

O

O

N

O

O

N

OO

N

OO

N

OO

N

OO

N

OO

Crosslinked UltraLinkpolymer containingazlactone groups

FIGURE 15.15 Ultralink resins are prepared by the copolymerization of N,N’-methylene bisacrylamide and the reactive monomer vinyl-dimethyl azlactone. The ratio of these two monomers used in the polymerization process controls the crosslinking and the degree of azlactone functionality in the final support.



N O

O

Azlactone group withinUltraLink matrix

HN

O

O–

Hydrolysis tocarboxylate

H2O

FIGURE 15.16 Azlactone groups can hydrolyze in aqueous solu-tion, which results in ring opening and the formation of a carboxylate group at the end of a short spacer.

N O

O

Azlactone group withinUltraLink matrix

+

Amine-containing ligand

H2N

HN

HN

O

Coupling via anamide bond linkage

FIGURE 15.17 An amine-containing ligand can react with an azlactone group by attack at the electrophilic carbon atom of the car-bonyl to result in ring opening and formation of an amide bond link-age with the ligand.

TABLE 15.1 UltraLink Swelling Properties in Various Solvents

Solvent Solubility Parameter (cal/ml) Volume (ml/gm) Relative Swelling2 (%)

Dry particles – – 4.5 0

Aqueous Solvents Water 23.4 8.10 100

1-N HCl – 8.08 99.4

1-N NaOH – 7.98 96.6

1-M NaCl – 7.90 94.4

Nonaqueous Solvents Dimethyl sulfoxide (DMSO) 12.0 8.50 111.1

Methanol 14.5 8.33 106.4

Formamide 17.2 8.25 104.2

Dimethyl formamide (DMF) 12.1 8.25 104.1

Ethanol 12.7 8.08 99.4

Acetonitrile 11.9 8.00 97.2

Isopropanol (IPA) 11.5 7.98 96.7

Acetone 9.9 7.82 92.2

Tetrahydrofuran (THF) 9.1 7.40 80.6

Methyl ethyl ketone (MEK) 9.3 7.07 71.4

2-Ethylhexanol 9.5 6.70 61.1

Ethyl acetate 9.1 6.50 55.6

Dichloromethane (DCM) 9.7 6.26 48.8

Methyl isobutyl ketone 8.4 5.5 27.8

Heptane 7.4 5.15 18.1

Triethylamine (TEA) 7.4 5.15 18.1

Decane 6.6 4.95 12.5

Toluene 8.9 4.95 12.5

Tetrachloroethylene (TCE) 9.3 4.85 9.7

2Relative percent swelling equals the amount of swelling in a solvent from dry beads relative to the amount of swelling in water = ([ml in solvent] – [ml of dry particles]) × 100/([ml in water] – [ml of dry particles]) = ([ml in solvent] – 4.5 ml) × 100/3.6 ml.



but the exact swelling factor is production-lot depen-dent, so the user should consult the lot-specific infor-mation before use. The amount of coupling reaction solution to use with a given amount of dried par-ticles should be equal to at least twice the amount of the hydrated volume of beads that is anticipated after swelling. This is due to the fact that an equal amount of solution to the hydrated bead volume will be com-pletely taken up within the pore volume of particles during the hydration process and a proportion of at least 50% slurry of gel in the reaction solution is needed to allow proper mixing during the reaction.

The dried UltraLink particles are supplied with a small amount of Triton X-100 coating within the par-ticles to facilitate rapid uptake of water into the pore structures without entrapment of air. For this rea-son, there will be an inherent absorbance at 280 nm to the reaction medium due to the release of detergent from the support. Unfortunately, this interference will prevent the determination of the amount of pro-tein coupled during the reaction simply by measuring the difference in absorbance at 280 nm of the solution before and after coupling, as can be done with many other coupling methods. Protein coupling can, however, be determined using the BCA protein assay reagent (Thermo Fisher) to measure the amount of immobi-lized protein directly on the surface of the beads after immobilization.

The handling of UltraLink supports can be per-formed using spatulas for manipulation, paddle stir-rers or rotators for mixing, and standard filter funnels with membranes or fritted glass filters for washing. The matrix is very robust and is not easily damaged in use. The methods used for the immobilization of proteins or other affinity ligands onto azlactone-activated sup-ports are described in subsequent sections within this chapter.

Toyopearl HW SupportsToyopearl supports are comprised of a synthetic

polymeric matrix that is commercially available from Tosoh Bioscience. The matrix is made from the copo-lymerization of glycidyl methacrylate, polyethylene glycol (PEG), and pentaerythritol dimethacrylate to form a complex structure that is hydrophilic and rich in hydroxyl groups for activation and coupling of affin-ity ligands (Figure 15.18). The polymerization of the two acrylate-containing monomers is combined with the covalent linkage of the PEG–OH groups to the gly-cidyl epoxides, thus forming pentaerythritol and PEG crosslinks within the porous polymer network mak-ing up the bead internal topography. The PEG chains are very hydrophilic and certainly contribute heavily to the biocompatible nature of the support, while the

pentaerythritol dimethacrylate crosslinks add rigidity by locking the polymer strands together.

Toyopearl-type supports have moderate mechani-cal stability and are able to tolerate pressures of at least 45 psi (3 bar) without bead collapse. This makes possible reasonable linear flow rates and large column configu-rations for process-scale separations or automated sepa-rations using an FPLC or HPLC system for smaller scale operations. A type of Toyopearl support containing a higher degree of crosslinking, and thus greater rigid-ity for higher pressure resistance, is available under the designation “PW,” but this matrix has lower capac-ity than the HW resins for protein purification applica-tions. The larger-particle-size versions of the support will be able to achieve higher linear flow rates at lower pressure drops across a column bed and thus be capa-ble of reach higher flow rates than the smaller particle grades. The matrix is also quite robust to handling, mix-ing, and chemical reactions carried out during activa-tion and coupling procedures. Toyopearl supports are able to withstand extremes in pH (2–12), high salt or buffer concentrations, the presence of denaturants, and a solvent exchange into nonaqueous conditions without suffering damage to the particles.

The wide variety of Toyopearl supports available commercially come pre-swollen as thick slurries in aqueous solution. A wide selection of different par-ticles sizes and porosities is available for different applications, depending on the resolution of separa-tion desired and the size of the column being used. Typically, smaller particles will be more appropriate for analytical separations, while larger particle diam-eters are more suitable for bench-scale to process-scale separations. The different exclusion limits available reflect the fact that the matrix was originally devel-oped for gel filtration or size exclusion chromatography and is still used for these applications. Small particle sizes of the support can be obtained that consist of 20 to 40-μm spherical beads and are designated “S” grade (for “superfine”), while medium particles are in the range of 40 to 90 μm (“M” grade), larger particles that have diameters of about 90 to 120 μm (designated “C” grade for “coarse”), and very large particles of diame-ters ranging from 100 to 300 μm (designated “EC” grade for “extra coarse”). The particle size distribution gen-erally gets broader as the particle grade and diameter increase, thus giving the smaller-particle-sized supports the potential to provide greater peak resolution in gel filtration chromatographic separations. In general, the larger particles provide less back pressure for a given flow rate and are more amenable to large-scale chro-matography, while the smaller particles provide greater resolution and are more appropriate for analytical sepa-rations where peak resolution is important.



For affinity chromatography use, the Toyopearl sup-port having an appropriate exclusion limit (correlating to its maximal pore size) may be chosen to best accom-modate the molecules being separated. For protein separations, typically the supports having the larger exclusion limits will provide the best opportunity for protein–ligand interactions in affinity separations, especially when two large proteins have to interact within the pores. For routine affinity purifications, the Toyopearl HW-65 or HW-75 resins will provide the best performance, as the exclusion limit of these gels are up

to 5000 kDa, with a fractionation range for size exclu-sion chromatography listed as approximately 50 to 5000 kDa. Pores of this size will allow even the largest macromolecules to interact within the matrix, including use in applications involving the capture of interacting proteins using co-immunoprecipitation procedures.

Toyopearl supports contain mainly secondary hydroxyl groups within its structure due to the presence of the pentaerythritol crosslinks as well as resulting from the creation of secondary hydroxyls upon epoxide group coupling with another hydroxyl within the web

Pentaerythritol dimethacrylateGlycidyl methacrylate Polyethylene glycol(PEG)

O O

O O

HO OHO

O

OH

OOH

+ +

O

O

O

O O

O O

HO OH

n

O

O

O

O

OH

On

O OHO

O O

O O

HO OH

OO

OHO

O

O

OH

OO

OH

n

O

O

O

O

HOOHO

O

OHO

O

O

O

O

HO

HO

O

O

O

O

HO

HOO

O

HO

OH

O

n

O

Complex, crosslinked Toyopearlcopolymer containing hydroxyl groups

and hydrophilic PEG chains

FIGURE 15.18 Toyopearl supports are constructed from the copolymerization of glycidyl methacrylate, the crosslinking monomer pen-taerythritol dimethacrylate, and polyethylene glycol (PEG). The epoxy monomer is also able to provide crosslinks within the support with neighboring hydroxyl groups and also incorporate PEG chains within the structure by reacting with the epoxides, which provides significant hydrophilicity. The support also contains an abundance of primary and secondary hydroxyl groups that can be used to couple affinity ligands.



of polymers. There is also some potential for primary hydroxyls being present if epoxide hydrolysis occurred without coupling to another hydroxyl group or some of the PEG–OH groups remained uncoupled at one end within the gel. Using any of these hydroxyls, the support can be activated through any of the activation reactions designed to create an electrophilic reactive group. This includes the use of such common activation reagents as CNBr, CDI, tresyl chloride, tosyl chloride, bis-epoxide, and divinylsulfone. However, the sup-port cannot be directly oxidized with sodium periodate to give aldehydes, as in the case of agarose-containing matrices; nevertheless, an intermediate modification with glycidol can be carried out to create polymer grafts on every hydroxyl within the gel that terminate in diols, which afterward can be oxidized to provide abundant aldehydes for immobilization through reductive amina-tion (similar to the procedure previously mentioned for Trisacryl supports). The initial glycidol derivatization process results in the small epoxide coupling to internal hydroxyls on the support and forming a terminal diol as the epoxide ring opens. Additional glycidol mole-cules then can react with these newly formed hydroxyls, thereby creating polymer grafts. Subsequent oxidation with periodate then creates an aldehyde at the end of each glycidol chain with simultaneous loss of one mol-ecule of formaldehyde. Coupling of amine-containing ligands on this derivative can be achieved through Schiff base formation followed by reductive amination to form a secondary amine linkage (see Section 2.1).

Separon HEMA SupportsCopolymer particles made of 2-hydroxyethyl meth-

acrylate have been used to create highly hydrophilic chromatography supports that have abundant pri-mary hydroxyls, which can be used for activation and coupling of affinity ligands. This synthetic support made from the copolymerization of HEMA and eth-ylene glycol dimethacrylate was developed decades ago in Czechoslovakia and commercialized by Tessek, Ltd., under the name Separon HEMA (Figure 15.19). The dimethacrylate crosslinking monomer adds rigid-ity to the HEMA polymer, and the ratio of crosslinker to HEMA in the final matrix controls the relative poros-ity of the support as well. A molecular model of a por-tion of the crosslinked HEMA polymer is shown in Figure 15.20, which illustrates the abundant hydroxyl groups present within the matrix. The particles are spherical and are available in sizes of 10-μm and 60-μm average particle diameters. HEMA supports also come in a range of porosities from 40 to 2000 kDa that provide abundant choices of porosity for size exclusion chroma-tography applications, and the degree of porosity can be chosen based upon the size of the molecules being separated. Particles having the largest pore size should be selected for general affinity chromatography sepa-rations using macromolecules, because the interior of the particles is able to better accommodate interactions between large biomolecules.

The high density of primary hydroxyl groups on the HEMA support allows convenient derivatization with

OH

OO

OH

O

O

OH

OO

HO

O

O

OH

OO

OH

OO

OH

O

O

OH

OO

HO

O

O

OH

O

O

OHOO

OHOO

HOO

O

OH

OO

HO

O

O

OH

O O

HO

O

O

HOO O

OH

O O

HOO

O

OH

O

O

OH

O

O

HO

O O

HOO O

OH

O O

OH

O

O

OH

O

O

OH

O

O

OO

O

O

Poly(hydroxyethyl methacrylate)(pHEMA)

FIGURE 15.19 Poly(hydroxyethyl methacrylate) (pHEMA) supports can be made from the copolymerization of 2-hydroxyethyl methacry-late (HEMA) and the crosslinking monomer ethylene dimethacrylate. The result is a support that is very hydrophilic and contains numerous primary hydroxyl groups, which can be activated for the coupling of affinity ligands.



typical activation reagents to form reactive, electrophilic intermediates capable of coupling affinity ligands. All of the common activation methods for hydroxyls described in this chapter may be used with HEMA, except for the use of periodate activation, which requires diols to form aldehydes, and this support does not contain any periodate-oxidizable hydroxyls.

Although HEMA-based supports contain esters from each hydroxyethyl group linked to the methacry-late monomers, the ester bonds are extremely stable due to the configuration containing a tertiary alpha-carbonyl ester. It does not easily hydrolyze nor does it react with nucleophiles such as amines. The base sup-port is stable to extremes of pH (2–12) and will tolerate NaOH sanitization procedures without breakdown of the polymeric structure. Two activated versions of the HEMA support are available from Tessek for the immo-bilization of affinity ligands: epoxy and vinyl sulfone. However, other activation methods may be designed from the base support for custom coupling purposes. In particular, glycidol modification may be carried out to create grafted polymers within the matrix that contain

terminal diols on each chain. As described previously in the section on Toyopearl supports, these diols can be periodate oxidized to form terminal aldehydes for use in reductive amination immobilization procedures (see the corresponding section under activation methods in this chapter).

UNOsphere SupportsAnother innovative polymeric chromatography

support is produced by Bio-Rad under the name UNOsphere. The base matrix construction is unique in that the beads are said to contain “through-pores” that are a minimum of 0.5 μm in diameter with virtu-ally no pores of less than 0.1 μm. This is reminiscent of Perceptive Biosystems’ HPLC supports that were devel-oped during the 1980s, which also are claimed to have larger through-pores, which make faster flow rates possible for analytical separations (Jungbauer, 1996; Gallant, 2004). The through-pores allow the mobile phase to enter into the internal spaces of the bead structure by convective flow rather than just by diffu-sion. In theory, this permits faster on and off rates for interactions with immobilized ligands than is normally possible using supports that only have convective flow around the beads (Figure 15.21). The majority of chromatography resins restrict internal bead access by diffusional processes originating from outside the par-ticles. In perfusion supports, the larger through-pores inside the particles allow convective fluid flow to reach sites deep within each bead, thus increasing the effec-tive volume of the support that can be used for affinity interactions during rapid chromatographic operations. The theory of intraparticle flow through perfusive sup-ports and monoliths has been studied in great detail (Hamaker and Ladisch, 1996).

The UNOsphere polymeric resin is made from the copolymerization of functional and nonfunctional monomers in addition to a portion consisting of cross-linking monomer to form the final matrix. The resultant

FIGURE 15.20 The three-dimen-sional, space-filling molecular model of a portion of a pHEMA support show-ing the numerous hydroxyethyl groups and a crosslink made from ethylene dimethacrylate.

Perfusive bead withlarge through-pores

Diffusive bead withsmaller pore structures

FIGURE 15.21 An illustration of the difference between a perfu-sive chromatography support containing large through-pores and a typical diffusive chromatography support which contains mainly dif-fusion-limited access to the inner pore structure of each bead.



beaded support has a particle size range of about 45 to 90 μm with a mean diameter of 60 μm. The spheri-cal particles form a chromatography support that is analogous in its internal pore structure to some of the monolithic-type supports that are formed by polymer-ization of monomers directly within a column to pro-duce a single continuous cylindrical matrix. It is also similar to the use of affinity membranes that contain a torturous pore structure with large and small pores where both convective flow and diffusion take place simultaneously throughout the matrix.

The large cavernous structures of through-pores within UNOsphere resins are constructed by careful selection of polymerization conditions, as described in U.S. patent 6,423,666 by Liao and Hjerten (2002). Just like most other polymeric supports, the particles are formed by suspension polymerization using a monofunctional monomer that may contain functional groups or reactive groups. For instance, the proper choice of monomer can yield a beaded chromatog-raphy resin containing ion-exchange functionalities, hydrophobic groups, or reactive derivatives able to spontaneously couple affinity ligands. One key to form-ing the large pore structure is controlled use of a cross-linking monomer during the polymerization reaction. Particularly, it has been found to be important to keep the concentration of crosslinker to a mole fraction of about 0.3 to 0.4 relative to the total monomer mixture. The construction of the through-pores is accomplished by the addition of some hydrophobic character to the monomers as well as the use of additives that modify the polarity of the aqueous medium during polym-erization. Additives such as sulfate or other lyotropic agents can be used in addition to the potential use of hydrophilic polymers such as PEG, dextran, or methyl cellulose.

The base support can be used in process purification applications that involve large column sizes or high linear flow rates (up to 600 cm/h) without particle col-lapse or encountering severe back pressures. The poly-meric support also can be sanitized using 0.1-N NaOH or regenerated after use with denaturants such as 6-M guanidine. A study of IgG purification on a UNOsphere support containing immobilized protein A was pub-lished by Perez-Almodovar and Carta (2009).

A reactive derivative of the UNOsphere support is available that contains epoxide groups, which can be used to immobilize affinity ligands containing amines, thiols, or hydroxyl groups (see the section on activation and coupling reactions, this chapter). It is supplied dry to ensure stability of the reactive group upon storage. The support contains approximately 50 to 132 μmoles/g of reactive epoxy groups, and upon addition to an aque-ous solution hydration will cause the matrix to swell to approximately 5.5 to 8.0 ml/g of dry resin. The dry,

activated support can be added directly to a ligand solution for coupling. The optimal pH range for cou-pling is 9 to 13 in a buffered solution containing at least 10-mM buffer salts to prevent pH changes during the reaction. Thiols can be coupled to the support at the lower pH, amines can be coupled in the middle of this range, and hydroxyl-containing ligands require pH 13 for efficient coupling. UNOsphere supports are not available in a hydroxylic form that could be used for custom activation and coupling of affinity ligands.

Eupergit SupportsCertain polymeric supports have found use for the

immobilization of enzymes, which are often used as immobilized reactors in the manufacture of foods, anti-biotics, and other organic compounds. Many of these supports are used in ton quantities annually for large-scale production processes. One such support that has found more use as an immobilized reactor than in affin-ity chromatography is Eupergit. These porous beads are made of methacrylate derivatives and were originally manufactured by Rohm Pharma Polymers, which was a unit within the Specialty Acrylics division of Degussa, AG in Darmstadt, Germany, but since 2007 this tech-nology became a part of Evonik Industries. Eupergit is made by a copolymerization of glycidyl methacrylate, allyl glycidyl ether, methacrylamide, and the cross-linking agent N,N’-methylene bis-methacrylamide. The Eupergit C support that contains reactive epoxide groups has become very common for use in the prepa-ration of immobilized reactors (Boller et al., 2002; Berrio et al., 2007) (see Chapter 1, Section 3.4, for a discussion of immobilized reactor applications). Unfortunately, after Rohm Pharma’s merger with Evonik Industries, the Eupergit support was discontinued; however, some distributors still sell small quantities of it for research purposes (Sigma Aldrich).

Sepabeads, ReliZyme, and ReliSorb SupportsAlthough Eupergit is no longer manufactured com-

mercially (see previous section), other polymeric sup-ports continue to be used to prepare immobilized reactors for production purposes. One such material is called Sepabeads and is produced by Resindion, which is a subsidiary of Mitsubishi Chemical. This support material consists of a porous polymer made entirely of polymethacrylate which is formed into spherical white opaque beads. Sepabeads is available in a num-ber of different formats including two different pre-activated supports containing epoxide groups ready to couple ligands, two amino functionalized supports containing different length spacer arms, a diol deriva-tive that can be periodate oxidized to create aldehydes for coupling amine-containing ligands, and two deriva-tives especially designed for hydrophobic interaction



chromatography: a C4 butyl and a C18 (Figure 15.22). The particle size of Sepabeads is available in two dif-ferent size ranges: an “S” grade with a particle size dis-tribution of 100 to 300 μm and an “M” grade having a particle size distribution of 200 to 500 μm.

The polymethacrylate Sepabeads support has an average pore diameter of approximately 10 to 20 nm, which is rather small for typical affinity chromatogra-phy resins. The average IgG antibody is about 11 nm in diameter, so large proteins may not be accessible to the smaller pore openings. However, a more recent itera-tion of the Sepabeads-type polymer was introduced under the name ReliZyme, and this material contains average pore sizes in the range of 40 to 60 nm, which are more amenable to protein chromatography applica-tions. All of the derivatives available for the Sepabeads products also are available in the larger particle ReliZyme beads.

Resindion’s Sepabeads and ReliZyme particles, how-ever, were developed with a view toward their use as immobilized reactors rather than for applications in affinity chromatography. Most enzyme reactors are prepared through adsorption or covalent immobiliza-tion of enzymes onto solid-phase supports, and their subsequent use is to act as catalysts in the transforma-tion of substrates into desired products. According to the company, an enzyme can be immobilized onto the Sepabeads or ReliZyme supports using one of four different approaches: (1) passive adsorption, (2) ionic interaction, (3) crosslinking, and (4) covalent coupling with a reactive group on the particles. Enzyme reactors have been prepared successfully using all of these strat-egies, although the most robust materials are formed using covalent coupling, which provides low leaching potential and a high degree of reusability.

A third resin material developed by Resindion is also potentially suitable for affinity chromatography

applications. The ReliSorb supports are polymethacry-late spherical particles available in three different particle size ranges: 50–150 μm, 75–200 μm, and 200–500 μm, with corresponding mean diameters of 90 μm, 120 μm, and 300 μm. The larger average particle size of the ReliSorb resins permits high-flow chromatography over a wide range of linear flow rates. The spherical particles are extremely rigid with low swelling char-acteristics when hydrated in aqueous solvent. In addi-tion, the pore structure of this support is in the range of 40 to 50 nm, thus making it useable for affinity chroma-tography separations using proteins and other biologi-cal macromolecules. The linear velocity of the largest particle size support can exceed 10 m/h with a pressure drop of only 0.5 bar/m of column bed height.

The ReliSorb particles also can be used in an expanded bed chromatography application, because the high specific gravity of the particles ( >1.1 g/ml) permits controlled bed expansion when used in a reverse-flow mode. A packed bed of resin will expand from 100% to 200% over the linear flow rate of 200 cm/h to 300 cm/h, when used in an expanded bed separation system. This important characteristic allows crude samples to be sep-arated on a bed of ReliSorb matrix without clogging the column. These properties are similar to the Streamline support materials from GE Healthcare.

The ReliSorb supports are available in a number of different derivatives including epoxide-activated, an iminodiacetic acid form for immobilized metal chelate chromatography, a diol derivative that can be periodate oxidized to form aldehyde groups, two amine function-alized resins, three different supports for hydropho-bic interaction chromatography including a phenyl, butyl, and a C18 support material, as well as a number of derivatives for ion-exchange chromatography. The epoxide- and amine-containing supports may be used for custom immobilization of affinity ligands containing amines, thiols, hydroxyls, and carboxylates. The diol-containing support may also be activated using a num-ber of different reaction strategies described later in this chapter (Section 2.1), which permits further customiza-tion of the support for particular applications.

Poly(Vinyl Alcohol) SupportsCrosslinked poly(vinyl alcohol) supports have been

used as resins in column-based separations, particu-larly involving high-performance liquid chromatog-raphy (HPLC) (Battinelli et al., 1996) (Figure 15.23). Poly(vinyl alcohol) (PVA) copolymers are typically made from the polymerization of poly(vinyl acetate) followed by the removal of the acetate ester by hydro-lysis to create the PVA derivative. Itagaki et al. (1989) described the production of porous PVA particles for use in chromatography applications. The beads are formed by dissolving the linear PVA polymer in an aqueous

Diol

Carboxylate Epoxide Primary amine

Iminodiacetic acid

O

OHOHN

O

O–

O–

O–

O

NH2O

FIGURE 15.22 The derivatives of Resindion chromatography supports that are available commercially.



solution containing a salt and dispersing the solution into a non-miscible organic solvent with stirring. The gelatinous polymer beads that form within the aqueous phase are stabilized through crosslinking with a bifunc-tional crosslinker, such as glutaraldehyde. One popular form of PVA copolymer is the Carbowax supports that are typically made from a combination of polyethyl-ene glycol (PEG), polydimethylsiloxane, and PVA (note that Carbowax is a trade mark of Dow Chemical, which refers to their line of products incorporating PEG com-pounds). These resins are often used for separating compounds in analytical gas chromatography or HPLC through differential interactions via hydrogen bond formation with the ether oxygens and hydroxyls mak-ing up the beaded matrix. Supports made of PVA are extremely hydrophilic due to the abundant –OH groups. Derivatives of these supports have been prepared through esterification or by activation of the hydroxyl followed by covalent linkage of affinity ligands. For instance, a reverse-phase HPLC support was prepared by esterification of dodecanoic acid onto the hydroxyl groups to form a C18 column able to separate com-pounds based on hydrophobic interactions (Battinelli et al., 1996).

Composite chromatography supports containing an inner core of a hydrophobic matrix and an outer shell of PVA have been created to make a biocompatible hydro-gel for protein separations (Leonard et al., 1995). A core particle made from porous polystyrene/divinyl benzene copolymer as modified through adsorption of PVA fol-lowed by stabilization by crosslinking. The resultant resin retained its ability to be used for size exclusion chromatography while the PVA outer layer provided hydrophilicity and decreased nonspecific binding toward proteins.

PVA polymers have also been used as carriers in the formation of drug conjugates. Kakinoki et al. (2008) cre-ated a paclitaxel–PVA conjugate to a linear 80-kDa PVA water soluble synthetic polymer to deliver therapeu-tic doses of the drug to tumor cells in vivo. The aque-ous solubility of paclitaxel was dramatically enhanced through conjugation to the extremely hydrophilic PVA.

Membrane SupportsThe use of porous membranes for the separation of

proteins and other biological materials has existed for decades and remains an important component for puri-fication, concentration, diafiltration, and removal of par-ticulates including viruses and bacteria contaminants. Membranes are used routinely in processes ranging from small research-scale operations to huge biopro-cess-scale production procedures in the pharmaceutical, biotechnology, and food industries. Synthetic and inor-ganic membrane supports are available with a broad range of porosity characteristics and structural composi-tions. Membranes may be constructed with large pores that permit macromolecules to freely pass through without being retarded or they may be made with smaller pore characteristics for selected filtration of mol-ecules above a certain molecular size (e.g., ultrafiltration membranes). The internal pore structure of polymeric membranes is formed by the torturous entanglement of molecular strands, which leaves gaps and winding pathways through its three dimensional structure.

At a microscopic scale, the internal structure of a membrane is similar to that of a macroscopic sponge. The inner pores can absorb an aqueous solution within its open spaces and allow flow of a mobile phase directly through them at much faster linear flow rates than is possible using beaded chromatography sup-ports. A packed volume of porous particles typically allows the mobile phase to freely move through it only by traveling around the beads, whereas the interior porous structure of the particles is accessed by diffu-sion. The process of diffusion is slower than perfusive flow and delays the interaction of molecules traveling in the mobile phase with the internal structure of the porous beads by the time needed for molecules within the mobile phase to diffuse into and out of the particles. By contrast, most of a membrane’s internal pore struc-ture is directly accessible by a flowing mobile phase with minimal need for diffusion to reach inner struc-tures. The dominance of convective flow through mem-branes with minimal dependence on diffusional mass transfer is a key advantage of membranes over packed-bed chromatography methods. In principle, this allows for much faster affinity interactions between the mobile phase and a ligand attached to the membrane’s surface than is possible using columns of porous particles.

However, membranes often have lower functional density than beaded supports, which can lead to a lower capacity for an affinity molecule to bind with a desired target. To overcome this potential deficiency, composite constructions of particles embedded within membranes have been made to combine the higher capacity of beaded chromatography supports with the faster flow rates possible with membrane supports.

O

O

n

O

O OH

n

Poly(vinyl acetate) Vinyl acetate Poly(vinyl alcohol)

OH

O

OH–

FIGURE 15.23 The reactions involved in the synthesis of a poly(vinyl alcohol) (PVA) support, which contains a huge density of hydroxyl groups throughout its structure.



Figure 15.24 shows the inner pore structure of a track-etched PET membrane (polyethylene terephthalate) containing large 1-μm pores and having nanoparticles covalently attached to the surfaces within it (Ulbricht, 2006). The particles provide significantly increased capacity for coupling enzymes used as reactors for chemical transformations, which occur in the mobile phase as it passes through the membrane.

Membranes are available having a broad selection of porosities ranging from those designed for ultrafil-tration of biomolecules to ones more appropriate for filtration of particulates or microbial contamination. Ultrafiltration membranes can be selected to have pore sizes that can allow the passage of molecules below a particular molecular weight range while generally excluding molecules above that point. The molecular weight cutoff value of membranes actually is an approx-imate figure, not an exact exclusion limit, but it provides useful information about what biomolecules potentially can access the interior porous structure of the support and pass through. To use membranes for affinity separa-tions, the selection of molecular weight cutoff must be one that is high enough to accommodate the intended molecules being targeted so they can bind an immobi-lized affinity ligand attached within the membrane’s structure. Thus, most membrane-based affinity sup-ports utilize membranes having large pore sizes in the micron range that allow proteins to pass through, for instance >0.2 μm. This cutoff range is essential to pro-vide internal space for protein–protein interactions to occur within the internal membrane structures while also allowing for rapid mobile phase flow.

Commercial polymeric membranes are available in a wide range of chemical compositions, including hydro-phobic or hydrophilic polymers, having functional

groups on the matrix available for chemical modifi-cation, containing anionic or cationic groups for ion-exchange separations, or having affinity ligands for capture of specific target molecules. Similar to beaded chromatography supports, polymer membranes can be made from naturally occurring polysaccharides or synthetic polymers. Some of the more popular poly-meric compositions consist of cellulose (regenerated), nitrocellulose, cellulose acetate, polyamide (e.g., nylon), polyimide, polyester, polyvinylidene difluoride (PVDF), polyacrylonitrile, polysulfone, polyethylene, polypro-pylene, poly(vinyl alcohol) (PVA, crosslinked), pHEMA (crosslinked), and a number of copolymers or compos-ites of these polymers (Ulbricht, 2006) (Figure 15.25).

Nitrocellulose and PVDF have long been popular for use with proteins, especially in western blotting proce-dures in which electrophoretically separated proteins are non-covalently transferred and adsorbed onto the membranes and probed immunochemically using anti-body conjugates. These membrane supports also can be used in affinity separations using non-covalently adsorbed proteins as capture ligands to selectively purify target molecules. However, to prepare more stable covalent attachment of affinity ligands it is neces-sary to have a functional group available on the mem-brane that can be activated and used to couple a protein or other affinity molecule. In this regard, membranes containing amines, carboxylates, or hydroxyl groups provide the greatest flexibility in the selection of activa-tion and coupling chemistries available to immobilize affinity ligands. Hydroxyl-containing membranes are particularly desirable for use in affinity separations. Unmodified cellulose (including regenerated cellulose) membranes in addition to PVA and pHEMA mem-branes or composites provide excellent hydrophilicity

FIGURE 15.24 An SEM picture of the internal structure of a porous PET membrane-particle composite construction. The nanoparticles are covalently attached to the membrane inner pore structure and provide increased capacity for affinity separations or enzyme reactor applica-tions. Reprinted with permission from Ulbricht, M. (2006) Advanced functional polymer membranes. Polymer 47(2006): 2217–2262; copyright © 2006 Elsevier.



and low nonspecific binding with abundant hydroxyls for coupling ligands. Composites containing naturally occurring hydroxylic polymers such as chitosan or dextran also provide abundant functionalities for acti-vation and coupling ligands. Another benefit of hydrox-ylic membranes is that they do not generate residual charge character from the remaining functional groups after coupling an affinity ligand, as may be the case with amine-containing or carboxylate-containing mem-branes, which could create considerable positive or neg-ative charges, respectively.

An important monomer type that has been used to prepare affinity membranes is glycidyl methacrylate, which contains an epoxide group that can be used to immobilize affinity ligands through available thiols, amines, or hydroxyl groups. Copolymers of glycidyl methacrylate can be prepared with other hydrophilic monomers to provide the membrane composition of choice. The terminal epoxy groups on poly(glycidyl

methacrylate) membranes also can be hydrolyzed by acid treatment to create terminal diols, which then can be periodate-modified to form aldehydes. The alde-hydes can subsequently be used for coupling amine-containing ligands by reductive amination (see the corresponding section in this chapter on reductive ami-nation and also Chapter 3, Section 5, and Chapter 4, Section 4). For instance, Akgöl et al. (2002) prepared a membrane consisting of poly(hydroxyethyl meth-acrylate-co-glycidyl methacrylate) that contained hydroxyl groups from the pHEMA component and epoxy groups from the polymerized glycidyl monomers (Figure 15.26). This membrane was used to couple the enzyme cholesterol oxidase for use as an immobilized reactor. In addition, Teke and Baysal (2007) used a base membrane consisting of nylon-6 (polyamide) and then grafted glycidyl methacrylate onto it using benzophe-none as an initiator to create a reactive epoxide-contain-ing membrane, which was used for coupling urease.

Nitrocellulose

OO OO

O

n

ON+O–

O

ON+–O

O

ON+–O

O

ON+

O–

OO

N+O–

O

ON+–O

OCellulose

OOHO

OH

OH

OO

O

n

HOOH

OH

H

HN

NH

O

On

Nylon 6

O

OO

On

Polyester

OH

n

Poly(vinyl alcohol) Poly(hydroxyethylmethacrylate)

nn

Polyethylene

OH

O O

n

Poly(glycidylmethacrylate)

O O

n

O

Polypropylene

FIGURE 15.25 The polymer structures of common membrane compositions.



Polymer grafting onto existing substrates has been an important route to the creation of reactive groups or functional groups on the surface of a non-function-alized membrane. Grafting onto a block copolymer substrate can introduce novel characteristics into a membrane that normally could not be created through the original membrane synthesis (Koguma et al., 2000). The process of grafting can create new poly-mer arms or tentacles coming off of the original mem-brane surface (Figure 15.27). For instance, the grafting of new polymer strands onto the internal structure of porous membranes has been used to increase the cou-pling capacity of membranes for immobilizing affin-ity ligands (Muller, 1990; van Reis and Zydney, 2007). Covalent grafting can be done via solution-phase polymerization of vinyl monomers in the presence of a membrane substrate and an initiator. Many routes to initiating this type of grafting reaction have been described and some of them can be used to graft syn-thetic polymers onto natural polymer surfaces (Jenkins and Hudson, 2001). Once the polymerization reaction is initiated, a new graft copolymer will be formed within the solution and can be attached to the existing mem-brane through any remaining vinyl groups left over from its initial synthesis. Most often, however, link-ing groups have to be created on the original block polymer surface to facilitate covalent attachment of the new polymer. Covalent graft copolymer modifica-tion of membranes and surfaces may use irradiation (i.e., using electron beam or Co60) to form radicals on the existing substrate, which can provide initiation sites for graft polymer growth (Gürsel et al., 2008). New poly-mer grafts may also be created by linking a polymer tentacle to a membrane using traditional conjugation chemistry. Gacal et al. (2006) used the Diels–Alder che-moselective ligation reaction (Chapter 17) to covalently attach PEG or poly(methyl methacrylate) polymers to a polystyrene substrate, thus creating novel surfaces with completely different properties from that of the original polymer. Noncovalent grafting also can be per-formed through the adsorptive interaction of a func-tional polymer with the existing membrane substrate, which instead of forming polymer arms, forms a layer

of the new polymer onto the initial membrane, in some cases filling the inner pores (Yang et al., 2011). This type of composite membrane is similar in construction to the “gel-in-a-shell” structure of agarose-filled ceramic par-ticles (described previously in this chapter).

Membrane-affinity supports have also been formed by the combination of porous beaded chromatography particles, which have been entrapped or grafted within the tangled polymer strands of a membrane (Borneman, 2006). This composite construction of porous beads within a membrane scaffold creates a chromatography support having the high potential convective flow rates characteristic of polymeric membranes plus the high

Hydroxyethylmethacrylate

Glycidylmethacrylate

OH

OO OO

O

+

m

OO

n

OOH

OO

Poly(hydroxyethyl methacrylate-co-glycidyl methacrylate)

FIGURE 15.26 The structure of a poly(hydroxyethyl methacrylate-co-glycidyl methacrylate) membrane.

Glycidylmethacrylate

OO

O

Polymer substrate Free radicalformation

Electronbeam

+

O

O

O

O

O

O

O

O

O

O

O

O

O

O

O

O

O

O

Poly(glycidyl methacrylate)graft polymer chain growingoff of original polymer fromfree radical initiation point

FIGURE 15.27 General illustration of grafting onto an existing polymer substrate. Many different polymer types can be grafted onto substrates to create new properties or add functional groups for cou-pling affinity ligands.



capacity and increased surface area present within the internal pores of particles. Particle-loaded nonwoven fibrous membranes were developed by 3M (Markell et al., 1994) and commercialized under the trade name Empore. These membranes contain either deriva-tized polystyrene particles or silica particles within an inert 10% polytetrafluoroethylene (PTFE) matrix and are used primarily for solid-phase extraction (SPE). Another fibrillated membrane/particle construct was made by incorporating 3M Emphaze beads (UltraLink, see previous section, this chapter) into a PTFE mem-brane, which provides azlactone-reactive groups for coupling affinity ligands (Haddad et al., 2010). A similar membrane-entrapped particulate technol-ogy was developed by Millipore and used to produce chromatography supports in pipette tips (Zip Tips) (Kopaciewicz et al., 2000). In addition, particle-embed-ded membranes have been used for the immobilization of enzymes to use as reactors, forming supports of high activity due to the capacity enhancement made possible by the entrapped particles (Ulbricht, 2006). An SEM picture of a nanoparticle-embedded membrane was shown previously in Figure 15.24 of this chapter. All of these constructs provide the ease of use and stabil-ity that are typical of membranes with the higher capac-ity of packed bed chromatography supports. Unlike packed beds, however, the particle-loaded membranes maintain better matrix stability by holding the particles in place and not being susceptible to bed disruption, which can lead to non-uniform flow characteristics.

Particle-loaded membranes have been used in affin-ity separations by immobilizing ligands on the embed-ded chromatography particles. Virtually any affinity separations that can be performed using beaded sup-ports can be accomplished using particle/membrane composites. For instance, immunoaffinity applications using immobilized antibodies have been used success-fully in SPE techniques to target and capture small anti-gens out of complex solutions, such as drugs in serum samples (Dombrowski et al., 1998; Delaunay et al., 2000).

Inorganic membranes also can be used for affin-ity separations. Membranes made of porous inorganic materials have been prepared from the same base compositions as beaded chromatography supports, including porous glass, silica, alumina, and ceramic materials. Many of these membranes are more rigid than those made from polymeric materials and can be molded during construction to fit the shape of a desired flow path or device. An exception to the usual rigid-ity of most inorganic membranes is glass fiber materi-als, which often are used in filtration applications and come in flexible disks of all sizes. Inorganic membranes may be surface modified to contain functional groups or reactive sites for coupling affinity ligands. The methods used to introduce sites for covalent coupling

on inorganic substrates often make use of functional silanes to form a polymeric network containing side chains, which terminate in the functional group or reac-tive group of choice. These reagents and their uses are described in Chapter 13.

Monolithic SupportsChromatographic monolithic supports are similar

in construction to membranes, but rather than polym-erizing monomers into a thin membranous sheet, the polymerization process forms a single continuous matrix that is contained within a three-dimensional shape, which acts as a mold. The final monolith thus can be constructed within a column shaped like a cyl-inder for standard chromatography purposes or formed within another type of mold that would result in a cus-tom matrix shape. Early pseudo monoliths were con-structed from stacked membrane sheets or from rolls of membranes, which formed larger beds of a discon-tinuous matrix. True monolithic supports, however, are constructed of a homogeneous porous matrix that takes on a desired shape dependent upon the mold in which it is contained. One of the first references to the name monolith with regard to a chromatography support was by Noel et al. (1993), who they prepared a continuous matrix of cellulose that took on the consistency of a sponge-like material.

However, explorations into the use of a continuous-phase matrix for chromatography separations occurred as early as the late 1960s and early 1970s, wherein uni-form separation media were prepared from such mate-rials as pHEMA, poly(ethyleneglycol methacrylate), foamed polyurethane, and other polymers for poten-tial use in separations (Kubin et al., 1967; Ross and Jefferson, 1970; Hileman et al., 1973). Since that time, a wide variety of molded polymers have been used to form monolithic porous polymers useful for all types of separations, including immobilized affinity ligand applications (Svec and Fréchet, 1999; Svec et al., 2003; Neff and Jungbauer, 2011; Shin et al., 2011; Arrua et al., 2012).

An example of a commercial monolith support is made by BIA Separations (Austria). The “Convective Interaction Media” produced by BIA is made in the form of disks and cylinders (Champagne et al., 2007). The composition of the polymer monolith consists of a homogeneous unit made of polymethacrylate deriva-tives (Jungbauer et al., 2008). The polymerization process produces large through-pores of 1.5-μm diam-eter with interconnecting pores throughout the support, which permits interactions between even the larg-est proteins or biological macromolecules within the media (Zmak et al., 2003; Etzel and Bund, 2011; Neff and Jungbauer, 2011). The BIA monolith supports are avail-able in popular affinity derivatives, including protein A,



protein G, and metal chelate. In addition, an epoxy-activated version is offered for direct immobilization of amine-containing affinity ligands as well as a car-boxy and an amino form for coupling of custom affinity molecules.

Protista International AB also commercialized a monolithic chromatography matrix that contains unusually large pore structures. This “supermacropo-rous” support was produced by the polymerization of monomers in aqueous solution within a column at temperatures that cause frozen sections to develop within the forming polymer (Arvidsson et al., 2002, 2003). The freezing of the aqueous regions causes very large pores to form consisting of cavernous spaces of 10- to 200-μm diameter throughout the “cryogel” monolith, thus creating a series of connected through-pores for rapid convective flow within the support (Arvidsson et al., 2002, 2003) (Figure 15.28). A vari-ety of monomers can be used to create the monolith, including derivatives containing ion-exchange groups, functional groups for modification, or reactive groups for immobilization of affinity ligands. A hydrophilic hydroxylic monolith was created using 2-hydroxy-ethyl methacrylate to form a cryogel containing abun-dant hydroxyl groups for subsequent activation and coupling (Savina et al., 2007). Other cryogel composi-tions have also been formed using other acrylamide or methacrylate monomers (Persson et al., 2004; Savina et al., 2006; Plieva et al., 2008).

The porous structure of the cryogel monolith even allows cell separations to be performed directly within the monolithic support using immobilized affinity ligands specific for certain cell types. For instance, Dainiak et al. (2005) used an IMAC support to sepa-rate different microbial cells by their ability to bind to the metal chelate ligands. Similarly, Kumar et al. (2003) used immobilized protein A to capture lymphocytes within a monolithic cryogel support, and Arvidsson et al. (2002) used immunoaffinity and anion-exchange

monolith supports to capture microbial cells (Figure 15.29).

Polymeric monolithic supports are chemically simi-lar to the corresponding membranes or spherical beads made from the same polymers—only the shape and size of the support are different. Therefore, the techniques used to activate and immobilize affinity ligands on a monolith’s polymer surface are nearly identical to those used for other support materials of equivalent chemical composition. For an illustration of the polymeric struc-tures of typical monolithic supports, see the previous overview of beaded chromatography supports in this chapter. The major difference between a monolith and a beaded support in the methods of ligand coupling is how the matrix is handled and washed during the activation and coupling reactions. Particulate supports can be manipulated as suspensions and washed after each reaction step using filtration devices, but a mono-lithic support is usually handled directly within the column or mold in which it was formed. For instance, if a column containing a porous monolith consisting of a pHEMA matrix is to be coupled with a protein affin-ity ligand, then each step in the immobilization process would be carried out by pumping the reagents through the column in a chromatography mode involving a series of sequential additions, incubations, and wash-ing steps. This chromatographic reaction scheme must be carried out carefully to ensure that all parts of the monolith are treated equally and end up with the same density of reactive groups and immobilized ligands. This is not a trivial point, because reactants are added in the mobile phase and are pumped through from one end of the column and out the other end. As the reac-tions occur, the concentration of the reactants change as the solution passes through the matrix. This is true even in radial flow chromatography operations where the mobile phase travels from the center of a cylindrical shaped monolith through the side walls. In general, as reactions take place within the monolithic support, the

FIGURE 15.28 SEM images of the supermacroporous structure of a Protista monolithic cryogel support. Reprinted with permission from Arvidsson, P., Plieva, F.M., Savina, I.N., Loxinsky, V.I., Fexby, S., Bulow, L., Galaev, I.Y., and Mattiasson, B. (2002) Chromatography of microbial cells using continuous supermacroporous affinity and ion-exchange columns. J. Chromatogr. 977: 27–38; copyright © 2002 Elsevier.



reactant concentration in the mobile phase will decrease as flow travels through the support and any byproducts of the reactions that are released into the solution will increase as the reagent front passes through.

Thus, the rates and yields of the activation and cou-pling reactions will decrease as the mobile phase flows through the monolith polymer. This could potentially result in dramatic differences in the amount of activa-tion and ligand coupling that occurs from the top of the monolithic column bed to the bottom, if consid-eration of this effect was not taken into account. To overcome this phenomenon, enough reactant solution must be passed through the matrix at each step in the activation and coupling process to fully saturate the monolithic support from top to bottom. One method of accomplishing this is to prepare a large excess of reagent solution and continually recirculate the solution through the column until the reaction period is com-plete. Alternatively, an excess of reactant solution may be passed through the monolith and then the support

allowed to incubate in the solution of the reactant for an extended time to fully drive the reaction to comple-tion. This process also can be used when coupling affinity ligands to an activated monolith, in which the support is gently mixed in the ligand coupling solu-tion after an initial amount of ligand has been passed through the column to saturate all the internal pore structures. This is the general strategy recommended by BIA Separations for immobilizing ligands on an epoxy-activated monolith made from poly(glycidyl methacrylate).

2. ACTIVATION AND COUPLING OF AFFINITY LIGANDS TO

CHROMATOGRAPHY SUPPORTS

The matrix and activation choices available for the preparation of immobilized affinity ligands are two of the most important considerations for creating an

(A) (B)

(C)

FIGURE 15.29 SEM images of supermacroporous monolith supports capturing E. coli cells using an anion-exchange group on the matrix (A and B) and lymphocytes from blood samples using immobilized protein A (C). Reprinted with permission from Arvidsson, P., Plieva, F.M., Savina, I.N., Loxinsky, V.I., Fexby, S., Bulow, L., Galaev, I.Y., and Mattiasson, B. (2002) Chromatography of microbial cells using continuous supermacroporous affinity and ion-exchange columns. J. Chromatogr. 977: 27–38, copyright © 2002 Elsevier; and Kumar, A., Plieva, F.M., Galaev, I.Y., and Mattiasson, B. (2003) Affinity fractionation of lymphocytes using a monolithic cryogel. J. Immunol. Meth. 283: 185–194, copyright © 2003 Elsevier.

6212. ACTIvATIon And CouPLIng of AffInITy LIgAndS To CHRomATogRAPHy SuPPoRTS


optimal affinity support. The selection of an affinity ligand is obviously equally important, but for this dis-cussion it is assumed that the target molecule to be cap-tured and the affinity ligand to be immobilized have already been identified by the user. Matching an affinity ligand with the best possible matrix material as well as choosing the optimal coupling strategy for immobiliz-ing the ligand are both critically important factors for ensuring high specificity and low nonspecific binding in the final chromatography support.

One of the first criteria in choosing an appropri-ate immobilization strategy is that the method used to activate and couple a ligand to a given support must be compatible with both the ligand and the matrix. In one regard, this means that the chemistry of activation must be chosen to be compatible in its reactivity with the type of functional groups present on the support material. For example, if the support is a polymer that contains abundant hydroxyl groups, which is a com-mon characteristic for many matrices (see previous sec-tion), then an activation method should be chosen that will yield an activated intermediate based upon the presence of an initial hydroxyl. In addition, the reactive intermediate created from activating the support must be appropriate for coupling to the available functional groups on the ligand to be immobilized. For instance, an electrophilic reactive group that spontaneously cou-ples to an amine may be an optimal choice for matrix activation when immobilizing an amine-containing affinity ligand, such as a protein.

Another important consideration is the reaction medium that is used to activate the chromatography support. Some activation methods require nonaque-ous conditions in pure organic solvents, which poten-tially may harm a support material by causing collapse or shrinkage of particles with consequential changes to internal pore structures. Determining if a desired support can be exchanged from an aqueous environ-ment into 100% solvent and back again without severe shrinkage or damage often is a prerequisite to deciding on an optimal activation strategy. For instance, a cross-linked agarose support can be washed into anhydrous acetone (or other water-miscible solvents) for activa-tion and taken back into aqueous coupling buffer for immobilization of a protein without irreversible dam-age occurring to the particles. However, a beaded poly-acrylamide matrix that was initially swollen in water will collapse by being exchanged into acetone, which will so severely constrict the pores within each particle that activation and coupling cannot occur efficiently throughout the internal regions of the support.

When choosing an activation method, consideration also should be made to residual functional groups left over after activation and coupling of an affin-ity ligand. Most activation chemistries alter the initial

characteristics of a matrix to form the intermediate reactive groups. If this activated intermediate success-fully couples with a ligand, then the desired covalent bond will form, which often replaces part of the reac-tive group with the coupled ligand. However, if ligand coupling does not occur at every reactive site on the support, then there will likely be reactive groups or hydrolysis products remaining after the reaction. The presence of such residual groups, if they are active, will still have coupling potential toward molecules in a sample when the affinity support is put to use. If any reactive groups remain after immobilizing an affin-ity ligand, the potential exists to covalently couple molecules that are not desired to be bound to the sup-port and thus create sites for nonspecific interactions. Additionally, if hydrolysis of reactive groups occurs at the same time as ligand coupling, then another func-tional group may be formed in the process which rep-resents the hydrolysis product. In some cases, this may generate charge characteristics, such as negatively charged carboxylates being left over after a reaction. Residual charged groups may cause nonspecific interac-tions with sample molecules independent of the desired affinity interactions. For these reasons, there is typically a blocking step associated with the ligand-coupling process that eliminates any residual reactive groups or blocks potential sites of nonspecific binding by capping them with a nonrelevant molecule. In some instances, the blocking of residual charged groups may not be possible after ligand coupling, because the ligand itself may have similar groups present and therefore would be modified in the process. In the end, the most impor-tant factor is to consider all of the reactions involved with support activation and ligand coupling to decide the best combination of reactions to produce an optimal affinity support.

Other important things to consider when choosing an activation method include the hazardous nature of the reagents required (personal safety and waste dis-posal issues), the reaction rate and yield when cou-pling a ligand, whether the reaction is reproducible and controllable under the chosen conditions, and if the process can be scaled up and performed in large batch sizes with similar efficiency to bench-scale experi-ments. Some activation and coupling methods work well in small scale, but are difficult to handle and con-trol in process-scale operations. For instance, cyano-gen bromide activation, despite its hazardous nature, is relatively simple to perform in a fume hood using milliliter quantities of a support; however, attempts to scale up this process to activate many liters of a support can result in significant problems related to environ-mental hazards and the difficulties in rapidly activat-ing, washing, and using the activated resin. Therefore, each prospective activation strategy should be carefully



considered as to its relative ease of use and scalability in production before deciding upon a method of choice for coupling a desired affinity ligand.

In addition, the quantity of the support to use in the activation and coupling reactions is dependent on how much target molecule ultimately needs to be captured when using the affinity support in a chromatographic separation. The amount of support to use is also gov-erned by how much affinity ligand is available for coupling. For abundant affinity ligands, an excess of affinity support can be prepared without significant cost considerations; however, for expensive ligands such as some primary antibodies or recombinant pro-teins, the ligand cost and availability may be the gov-erning factors in deciding how much of an affinity support to prepare. Most of the protocols in this section involve the activation of from 1 ml to 100 ml of a sup-port material. The amount of material actually used to produce a desired immobilized ligand may be var-ied from the given protocols simply by proportionally adjusting the reagents used in the reactions.

If a protein is to be immobilized onto a matrix, ide-ally it should be dissolved in the coupling buffer at concentrations of at least in the low mg/ml levels. Many proteins to be used as affinity ligands perform well if the immobilization reaction is performed at con-centrations of about 3 to 6 mg/ml. In some cases, when maximal binding capacity is desired for the affinity support to capture a target molecule, a protein ligand may be coupled at concentrations much higher than this range, even >10 mg/ml. Most coupling reactions usually provide high yields of protein immobilization when using concentrations of up to about 20 mg/ml during the immobilization reaction. Immunoglobulin-binding proteins such as protein A are typically cou-pled to chromatography supports at the high end of this concentration range ( >6 mg/ml) to maximize the binding capacity of the resultant affinity support for purifying antibodies.

However, regardless of the type of affinity support being prepared, it is recommended that experimental optimization be done for the coupling ligand concen-tration and the reaction conditions to provide the best possible affinity support performance for binding a desired target molecule. If some proteins are immo-bilized at too high a density on the support, then non-specific binding may occur toward other molecules in a sample, or alternatively it may result in difficulty in eluting a bound target molecule due to extremely high affinity or avidity effects. Conversely, if too low a quan-tity of protein ligand is immobilized, the capacity and efficiency of binding to the desired target molecule may be unacceptably low. Of course, if only a small amount of a ligand is available for coupling, then using reaction concentrations down to the μg/ml levels may be carried

out with success, but along with the realization that the resultant binding capacity of the support will be lower as well.

If a small amine-containing ligand is to be immobi-lized on the support, its concentration in the coupling buffer should be at least three times greater than the level of active groups on the matrix to obtain decent yields. Many activated supports may contain about 20 to 40 μmoles/ml gel of reactive groups; therefore a small amine-containing ligand might be reacted at a level of 60 to 120 μmoles/ml gel. Higher concentrations will result in higher yields of coupling, but optimiza-tion of the resultant ligand density should be carried out to obtain the best performance of the affinity sup-port in its intended application.

A slightly different situation results if a diamine spacer molecule is to be coupled to an activated support to result in a free terminal amine. To avoid the reaction of the support with both ends of the linker the molecule should be dissolved in the coupling buffer at very high concentration. For instance, a 1.0- to 1.5-M concentra-tion of ethylene diamine will maximize coupling yield while preventing both ends of the compound from cou-pling to the support.

It is interesting to note that when coupling small amine-containing ligands or larger proteins to mic-roparticles or nanoparticles (see Chapter 14) the most frequently used immobilization technique involves activation of carboxylates on the particle surface using the carbodiimide EDC (with or without the addition of NHS or sulfo-NHS), which is then immediately used to react with the ligand. However, for larger particles used in chromatographic separations, the carbodiimide reaction is typically applied only when immobilizing small amine-containing molecules or diamine spacers, and then only in cases in which the molecules do not also contain carboxylates and thus have no potential for polymerization during the reaction. Most coupling methods involving chromatographic supports make use of a much wider variety of activation and coupling reactions to facilitate the immobilization of affinity ligands than those typically used with microparticles or nanoparticles. This statement does not mean that the methods described in this chapter cannot be applied to small particles—indeed they can with the appropriate changes to accommodate the differences in handling and washing, which typically involve centrifugation or magnetic separations as opposed to simple bulk filtra-tion as can be done with chromatography supports. The reactions discussed in this chapter can be applied suc-cessfully to any immobilization requirement, whether the solid phase is a planar surface, a microparticle, a nanoparticle, a membrane, or a larger porous bead.

This section describes the activation meth-ods commonly used to create reactive groups on



chromatography support materials. All of these reac-tions have been used successfully for many years and have abundant publications that describe the coupling of specific affinity ligands of all types. This section also contains suggested ligand coupling protocols following each activation procedure. All of these reactions should be performed with caution. Care should be taken to use the appropriate personal protective equipment to guard against exposure to potentially toxic compounds or flammable solvents. Most activation methods should be carried out in a fume hood wearing gloves and a lab coat as a minimum. Occasionally, a full-face shield or an organic mask also should be used to prevent chemical exposure to hazardous compounds.

2.1. Amine-Reactive Immobilization Methods

Periodate Oxidation and Reductive AminationOne of the most popular methods of immobilizing

affinity ligands on chromatography supports involves the formation of aldehyde groups on the matrix fol-lowed by a reductive amination coupling reaction to covalently link amine-containing molecules. Sodium meta-periodate is an oxidizing agent that can be used to create aldehydes on chromatography supports that contain diols. Hydroxyl groups present on adjacent car-bon atoms are a common feature in many resins made from polysaccharides, such as beaded agarose, cellu-lose supports, or cellulosic membranes. Diols may also be formed on synthetic polymeric support materials by careful choice of the monomers used during manu-facture or from the selected chemical modification of a support after production. For instance, glycidyl meth-acrylate monomers can be used to prepare chromatog-raphy matrices that will yield epoxide groups in the final beaded support or membrane. Hydrolysis of the epoxides under acidic conditions will generate termi-nal diols on the carbon atoms that originally made up the epoxide rings. In addition, the crosslinking of some support materials with epoxide-containing agents, such as the use of epichlorohydrin or 1,4-butanediol diglycidyl ether to stabilize and strengthen beaded agarose supports, may create additional periodate-oxi-dizable diols from leftover, hydrolyzed epoxide groups (Figure 15.30).

Diols can even be formed on support materials con-taining non-periodate-oxidizable hydroxyl groups through modification with the small epoxide-contain-ing compound glycidol (Hoyer and Shainoff, 1980; Shainoff, 1980; Guisán, 1988; Guisán et al., 1997). The epoxy group of this compound reacts with hydroxyl groups under alkaline conditions to form an ether bond with the associated formation of a terminal diol (Figure 15.31). Reaction of glycidol in excess with hydroxylic

supports results in the polymerization of the com-pound within the matrix through continued reaction with the hydroxyls produced as the epoxide couples. This may form branched polyglycerol chains in which each branch terminates in a diol, the formation of which dramatically increases the hydrophilicity of the matrix while forming the requisite periodate oxidiz-able components to create aldehydes for coupling affin-ity ligands. The amount of glycidol used in this process determines the ultimate size of the polymers formed and the potential affect on the internal porosity of the support, which can be significantly reduced even in highly porous gels (Eriksson, 1987). Glycidol modifica-tion has been used to immobilize and stabilize enzymes through multipoint attachment to the matrix, and the level of modification and subsequent activation to aldehydes can be controlled to produce between about 7 and 70 μmoles of reactive groups/ml gel (Suh et al., 2003; Tardioli et al., 2011; Nwagu et al., 2012).

Periodate oxidation of diols proceeds by cleavage of the associated carbon–carbon bond with concomitant oxidation of the hydroxyl groups to aldehydes. If the diol consists of secondary hydroxyls present within the sugar rings of a polysaccharide polymer, it will result in two aldehydes being formed per oxidation event along with scission of the ring; however, if the diols are terminal—in other words, containing one primary hydroxyl group adjacent to a secondary hydroxyl, such as those created from epoxide hydrolysis or glycidol modification—an oxidation reaction would yield one molecule of formaldehyde (released) and one aldehyde remaining on the support at every diol site. In addi-tion, if a series of three hydroxyls occur on neighboring carbons, such as in glucose-containing polymers like dextran, then treatment with periodate would result in the formation of one molecule of formaldehyde (from oxidation and loss of the central carbon atom) and two aldehydes generated from oxidation and cleavage of the glucose ring (Figure 15.32). The oxidation of polysac-charide polymers can be highly complex and has been studied extensively (Rinaudo, 2010). Periodate-oxidized supports prepared in this manner can be washed free of excess periodate (and formaldehyde) and stored for long periods without loss of aldehyde functionality.

Aldehydes (or ketones) can react with amine groups in aqueous solution to create reversible Schiff base linkages, the formation of which is enhanced at alka-line pH values. Schiff base formation is a dehydra-tion reaction that can reversibly undergo hydrolysis in aqueous solution to reform the carbonyl group and the amine. However, Schiff bases can be stabilized and made essentially permanent by reduction to a second-ary amine bond (Figure 15.33). Reducing agents that are suitable for this process include sodium cyanoborohy-dride and certain borane compounds (Borch et al., 1971;



Jentoft and Dearborn, 1979; Baxter and Reitz, 2002). Stults et al. (1989) showed that up to 50 mg of BSA could be immobilized per gram of a crosslinked agarose-beaded support using pyridine borane as the reduc-ing agent. Sodium borohydride may also be used to reduce a Schiff base, but it is about 5-fold stronger as a reducing agent than cyanoborohydride, and thus it could also reduce the aldehyde reactive groups used to create the initial Schiff base and it could reduce and cleave disulfides within protein molecules (Peng et al., 1987). Cyanoborohydride will not normally appreciably

reduce carbonyls, except under acidic conditions at pH 3 to 4 (Lane, 1972). Therefore, under slightly more alkaline conditions (pH 6–8), cyanoborohydride can be used to help drive the immobilization reaction to com-pletion—reducing the Schiff base interactions as they form—whereas stronger reductants may also reduce one of the starting reactants, thus potentially reducing yields.

It has been shown that the coupling efficiency and the resultant stability of the reductive amination pro-cess for immobilizing proteins on agarose supports

OO

OH

OO

HO

OH

O

O

OH

HO

O

OH

O

OH

HO

O

OH

O

Agarose

OOHO

OH

OH

OO

O

n

HOOH

OH

H

Cellulose

NH

O

OH

O

OHOH

O

OHO

OH

O

OHOHO O

OHOH

O

OHOH

O

Glycidol-modifiedsupports

O O

n

HO

HO

Hydrolyzed epoxy-containing supports

OOO

OH

OHO

OH

O

OH

OHHO

OHOHO

OH

O

n

O

HOHO

OHO

O

Dextran polymer

FIGURE 15.30 Polymeric supports and modified supports that contain hydroxyl groups suitable for activation and coupling of affinity ligands.



are both excellent (Domen et al., 1990). Proteins hav-ing abundant amino groups for immobilization typi-cally have high coupling yields, which can exceed 95%. Yields can be enhanced by forming the initial Schiff base in a reaction medium having a more alkaline pH of 9 to 10 and then reducing the pH toward neutral-ity for subsequent reduction of the iminium double bond (Hornsey et al., 1986). In addition, Ruzicka et al. (2006) showed that reductive amination reactions can be complete in just several minutes instead of the typi-cal recommended reaction times of hours. They also contended that the reduction step with cyanoborohy-dride is unnecessary, as protein ligands immobilized via Schiff base formation at high pH appeared to have little ligand leakage even when treated with 0.1-M HCl. Despite these results on microliter quantities of sup-port materials, care should be taken when coupling using much larger volumes of supports to determine the optimal reaction time for each step in the coupling process. Mixing and diffusion rates within bead slurries of quantities from hundreds of milliliters to multiple liters of gel can be quite different from reactions carried out in a micro flow cell with microliter amounts of gel.

In addition, while Schiff base interactions can be more secure due to increased stability gained from multipoint attachment with proteins—given that the probability of all the associated Schiff base linkages hydrolyzing at once and releasing a protein molecule is low—it is still highly recommended that stabilization of these linkages be done through reduction to prevent the possibility of hydrolytic leakage of ligand over time. In particular, if an affinity ligand is immobilized through only a single amine to form an iminium Schiff base group, then fail-ure to reduce that bond will result in significant ligand leakage over time by hydrolysis.

The following protocols involve various strategies to form the reactive aldehyde groups on support materials as well as the suggested coupling reactions for immobi-lizing amine-containing ligands. Note: The use of sodium cyanoborohydride is potentially toxic and dangerous. All operations should be performed in a fume hood while wearing appropriate personal protective equipment.

Periodate Activation ProtocolThis protocol is designed to create aldehyde groups

on diol-containing chromatography supports through

Supportcontaining

hydroxyl groups

+

OHO

Glycidol

OHO

O OHO

HO

O

O

O

OHHO

O

OHO

OH

O OHO

O OHOH

O

OHHO

O

HO OH

O

HO

O

OHOH

O

HO

HO

O

Branched glyceryl polymercontaining terminal diols

OH

OH

FIGURE 15.31 Glycidol reacts with hydroxyl-containing supports in a ring-opening process to create ether linkages, which in turn create additional hydroxyl groups. Glycidol can polymerize within the matrix to form branched polymers wherein each branch terminates in a diol, which may be periodate oxidized to form aldehyde groups.



oxidation with sodium periodate. This is useful for the activation of crosslinked agarose supports, cellulose- or dextran-containing beaded supports or membranes, and matrices that have been modified with glycidol or con-tain a hydrolyzed epoxide to create diol groups. Changes to the quantity of support material used from that described here may be done with corresponding propor-tional changes to the amounts of other reagents used.

1. Wash thoroughly 100 ml of a diol-containing chromatography support material (i.e., crosslinked agarose, dextran, cellulose, or a glycidol-modified support material) with at least several bed volumes of deionized water to remove any preservatives or other contaminants. The use of a sintered glass filter funnel works well for this purpose. Drain the support to a wet cake.

OHO

HOOH

O

OHO

HOOH

O

OHO

HOOH

O

OHO

HOO

O

O

Dextran containingD-glucose residues

NaIO4

NaIO4

NaIO4

O

O

4H2CO

OO

OO

O

O

O

OO

O

O

OO

OO

+

Polyether chaincontaining aldehdyes

OHO

O

OHHO

O

OH

OHO

O

O

HO

HO

OH

O

OHO

OH

OH

O

O

HO

HO

OH

OO

O

OH

OH

Cellulose polymercontaining D-glucose

residues

O

OO

O

OH

OO

O

O

OH

OO

O

O

OH

O

OO

O

O

OH

Polyether chaincontaining hydroxyls

and aldehdyes

Glycidol-modifiedsupport

OHO

OH

O

O

H

OO

Glyceryl polymerwith terminal aldehydes

2H2CO

+

4H2CO+

HO

FIGURE 15.32 Diol groups in support materials may be oxidized with sodium periodate to form aldehydes, which can be used to immobi-lize amine-containing ligands by reductive amination. Saccharide residues containing diols, such as within dextran or cellulose polymers, will undergo ring cleavage and opening along with the formation of two aldehyde groups per sugar. Glycidol modified supports will undergo oxi-dation with the formation of a single aldehyde at the end of each branched-chain polymer.



2. Dissolve 4.28 g of sodium periodate (NaIO4; MW = 213.89 g/mol) in 100 ml of water to prepare a 0.2-M solution. Mix the washed gel cake into the periodate solution to create a uniform suspension of the particles. Notes: At this level of periodate addition, the method will create a maximal level of aldehyde groups within the support as the internal diols are oxidized. If lower levels of aldehyde production are desired, then the amount of periodate addition can be controlled to provide this result. For instance, Guisán et al. (1997) determined that to create a support containing about 5 μmoles aldehydes/ml gel from a glycidol modified crosslinked agarose support (see the protocols which follow for glycidol modification), one could add a more dilute concentration of periodate to the modified support. To achieve similar moderated activation levels in this protocol, instead of using the recommended levels of periodate in step 2, dilute 5 ml of 0.1-M sodium periodate solution with 995 ml of water and then add this solution to the 100 ml of wet gel cake prepared in step 1. Conversely, to form about 75 μmoles aldehydes/ml gel on a glycidol modified agarose support, add 175 ml of 0.1-M sodium periodate to 825 ml of water and then add this solution to the wet gel cake. Stir these solutions for 60 min (for low-level activation) or 90 min (for higher activation) at room temperature to oxidize the supports to the desired level of activation, and then wash as in step 4. Another strategy that may control the level of aldehydes formed on a support is to regulate the initial

glycidol modification reaction to control the amount of diols formed (Guisán, 1988).

3. Continue the reaction for 90 min at room temperature with constant mixing using an overhead paddle stirrer or a rotating mixer. Do not use a magnetic stir bar, as this will damage beaded supports. If oxidizing a membrane, the support material should be gently agitated throughout the reaction in a bath of the periodate solution. Periodate oxidation takes place rapidly to form aldehydes within the matrix, and such reactions on a small scale may be complete in as little as 10 to 15 min. For larger quantities of gel, longer reaction times may be appropriate to ensure complete mixing of the oxidant throughout the support. For some resins, such as non-crosslinked agarose or cellulose, extended oxidation may cause damage to the structure of the particles or may actually dissolve them. For such supports, a very brief 5- to 10-min exposure to periodate may aid in maintaining the matrix structure while still forming sufficient aldehyde groups for immobilization of ligands. For extremely sensitive supports, scale back both the concentration of periodate and the time of reaction.

4. Wash the oxidized support with at least 10 bed volumes of deionized water to stop the reaction and remove excess periodate and any formaldehyde that may have been formed during the reaction. The aldehyde groups created on the support by this process are stable in aqueous solution indefinitely if a preservative is added to the slurry to prevent microbial growth. Store the activated gel until use at 4°C as a 50% (v/v) slurry containing a preservative.

Aldehyde-containing support

NaCNBH3

O

HH2N

Amine-containing ligand

+N

H

H2O

HN

Secondary aminebond formation

Schiff baseformation

FIGURE 15.33 Aldehyde-containing supports can be reacted with amine-containing ligands in the presence of a reducing agent such as sodium cyanoborohydride to couple the ligand through a secondary amine bond.



Use of Glycidol to Form Periodate-Oxidizable DiolsThe following protocols can be used to modify pri-

mary or secondary hydroxyl groups on support mate-rials to create diols susceptible to periodate oxidation. They can be used to modify supports that do not natively contain periodate-oxidizable diols, which are necessary to form aldehydes using oxidation. Polymeric support materials that can be used with success in this process include Trisacryl resins (containing primary hydroxyls from Tris-containing monomers) and Toyopearl resins (containing hydroxyls from pentaerythritol-containing monomers). In addition, other primary or secondary hydroxyl-containing supports such as those made from HEMA, PVA, and ethylene glycol methacrylate (EGMA) are candidates for modification with glycidol to create periodate oxidizable diols. Glycidol may also be used to modify amine-containing supports in a similar man-ner. The benefits of using hydrophilic modifications with glycidol to form polyglycerin spacers within a matrix for the coupling of affinity ligands have been documented (Rhemrev-Boom, 2009). Hyperbranched polyglycerols made from controlled polymerization of glycidol can form dendritic molecules as well as nanoparticles or microparticles containing pendent diols, which can be activated by periodate to form aldehydes for coupling ligands (Sunder et al., 1999, 2000).

Glycidol has also been used to modify crosslinked agarose despite the fact that agarose already has peri-odate-oxidizable diols within its matrix structure. In this case, glycidol forms branched polyglycerin tethers that project off the agarose resin and are highly mobile within the porous structure of each particle. Periodate oxidation of the terminal diols on these polyglycerin groups forms aldehydes that can be used to couple enzymes at multi-ple lysine amino sites on their surface. Such multipoint attachment aids in enzyme stabilization to create immo-bilized reactors of high thermal stability and reusability for industrial processes (Ichikawa et al., 2002; Ming et al., 2006; Kuroiwa et al., 2008). Site-directed mutagenesis of penicillin G acylase followed by immobilization on glyc-idyl-agarose resulted in increased stability and activity of the enzyme (Abian et al., 2004). This technique also can be used on hydroxyl-containing monolithic supports, membranes, and on the surfaces of hydroxylic microflu-idic channels to form reactive tethers for enzyme or affin-ity ligand immobilization (Albrecht et al., 2010).

The following methods should be carried out in a fume hood using the appropriate personal protective equipment to avoid exposure to the epoxide-containing reactant, glycidol. The first two procedures describe aqueous reactions of glycidol with hydroxylic supports in the presence of base, while the second one involves a non-aqueous reaction using BF3 etherate as catalyst for the epoxide reaction in dioxane.

(A) AQUEOUS GLYCIDOL REACTION FOR HIGH DENSITY MODIFICATIONS

The following protocol is designed to create a high concentration of glycidol polymer modifications throughout the hydroxylic support material. This will form numerous terminal diols for subsequent periodate oxidation, but the protocol also will create considerable polymer occlusion within the pore structure of some sup-ports. This procedure has been determined to work well for certain polymeric supports, but the optimal level of glycidol modification for a specific application should be determined experimentally. See method (B) for a reaction that will yield a lower degree of glycidol modification.

1. Wash 100 ml of a support material, for instance Trisacryl GF-2000 or Toyopearl HW-65F, with 500 to 1000 ml of deionized water using a sintered glass filter funnel to remove the preservatives or storage solutions that are present in the commercial products. Finally, wash the gel with 100 ml of 1-N NaOH (caution: caustic solution) and drain to a wet cake, but do not allow it to dry.

2. Carefully add the washed gel to 100 ml of 1-N NaOH and resuspend it with stirring.

3. Add 100 ml of glycidol solution to the stirring gel suspension along with 1 g of sodium borohydride (NaBH4). Note: With some matrices optimization of the amount of glycidol to add should be done to prevent unacceptable decreases in porosity due to internal glycidol polymer formation within the particles.

4. Continue the reaction overnight with constant stirring using an overhead paddle stirrer.

5. When the reaction is complete, carefully transfer the gel slurry to a sintered glass filter funnel, and wash extensively with water (1–2 l), 1-M NaCl (1 l), and again with water. The glycidol-modified gel can be stored until use at 4°C as a 50% aqueous slurry containing a preservative or oxidized immediately with periodate to form aldehyde groups according to the previous protocol.

(B) AQUEOUS GLYCIDOL REACTION FOR MEDIUM-DENSITY MODIFICATIONS

The following protocol will result in glycidol modifi-cations within a hydroxylic support material at a much lower level than method (A) above. This process has been found to work well for the modification of aga-rose gels to create numerous periodate-oxidizable sites within the matrix and to form a more hydrophilic inte-rior within the support (Guisan, 1988).

1. Wash 100 ml of a support material containing hydroxyl groups—for instance, agarose, Trisacryl



GF-2000, or Toyopearl HW-65F—with 500 to 1000 ml of deionized water using a sintered glass filter funnel to remove the preservatives or storage solutions, which usually are present in the commercial products. Finally, wash the gel with 100 ml of 0.375-N NaOH and drain to a wet cake, but do not allow it to dry.

2. Carefully add the washed gel to 100 ml of 0.375-N NaOH and resuspend it with stirring.

3. Very slowly add 34 ml of glycidol solution to the stirring gel suspension along with 1 g of sodium borohydride (NaBH4). The glycidol should be added dropwise to prevent the solution from heating beyond about 25°C.

4. Continue the reaction overnight with constant stirring using an overhead paddle stirrer.

5. When the reaction is complete, carefully transfer the gel slurry to a sintered glass filter funnel, and wash extensively with water (1–2 l), 1-M NaCl (1 l), and again with water. The glycidol-modified gel can be stored until use at 4°C as a 50% aqueous slurry containing a preservative or oxidized immediately with periodate to form aldehyde groups according to the previous protocol.

(C) GLYCIDOL REACTION IN DIOXANE

An alternative glycidol modification method was described by Eriksson (1987), in which a high-percent-age agarose gel that had been crosslinked with divinyl sulfone was further modified with glycidol in diox-ane. This process results in a more hydrophilic agarose matrix when supports containing a high percentage of agarose are used (up to 20% agarose).

1. Wash 10 ml of a support material with 100 ml of deionized water to remove preservatives and other storage solution components. Next, sequentially wash the support into dioxane by using slowly increasing concentrations of the solvent in water until the agarose is in 100% solvent. This can be done by washing first with 50 ml of dioxane:water at a ratio of 1: 3 (v/v), then washing with 50 ml of dioxane:water at 1:1, then washing with dioxane : water at 3 :1, and finally washing with 100 ml of 100% dioxane to fully remove the last traces of water.

2. Suspend the agarose support in an equal volume of dioxane and stir in a fume hood using a paddle stirrer or a rotating mixer. Add 3 ml of glycidol with mixing followed by 0.25 ml BF3 etherate.

3. React for 1 h at room temperature with continuous mixing.

4. When the reaction is complete, carefully transfer the gel slurry to a sintered glass filter funnel and

wash with 100 ml dioxane followed by a sequential transfer back into aqueous solution by reversing the dioxane:water ratios used in step 1. Finally, wash with 100 ml of water and store the gel at 4°C in an equal volume of water containing a preservative. The glycidol modified support may be periodate-oxidized to contain aldehyde groups according to the protocol described previously.

Immobilization of Ligands Using Reductive Amination

The following protocols are generalized procedures for the immobilization of amine-containing small ligands, spacer molecules, or proteins onto periodate-oxidized support materials, which contain aldehyde residues available for coupling by reductive amination. The reactions should be performed in a fume hood to avoid exposure to hazardous or toxic compounds, espe-cially the cyanoborohydride used in the reduction step. Two methods are described for reductive amination: the first one using a reaction buffer at physiological pH and the second protocol using an initial incubation at pH 10 to form the Schiff bases followed by reduction at pH 7.2. The first method works well for the major-ity of ligands and should be used for proteins that are sensitive to the more alkaline pH reaction. However, if ligand coupling densities or coupling yields are lower than desired, especially using proteins, the pH 10 pro-tocol typically provides increased coupling efficiency, which can exceed 95% yield.

The sodium cyanoborohydride used for the follow-ing reactions may be obtained as the pure solid or as a 5-M NaCNBH3 solution in 1-N NaOH (Aldrich). The solution is more convenient to use, because the solid may be susceptible to static charge during weighing, which causes fine particles to fly away and increases the hazards of handling the compound. The only caveat in using the alkaline cyanoborohydride solution is that after addition the final pH of the reaction should be checked and adjusted back to pH 7.2 if necessary.

(A) COUPLING AT pH 7.2

1. Wash 100 ml of the periodate-oxidized support material containing aldehyde groups into coupling buffer (0.1-M sodium phosphate, pH 7.2) and drain to a wet cake. Other buffer components may be added to the coupling buffer, such as the addition of 0.15-M NaCl or alternative buffer salts; however, avoid amine-containing compounds like Tris, glycine, or imidazole, which will compete in the reaction. In general, avoid buffer compounds containing primary or secondary amines as well as any other nucleophile that could react with an aldehyde. In addition, certain detergents and other amphiphilic components



have been shown to decrease the stability of the intermediate Schiff base and these additives should be used only after validating that they have no affect on the immobilization reaction (Viguera et al., 1990). Reductive amination coupling in aqueous solution has been shown to effectively occur between pH 4 and 10, with an optimal range for cyanoborohydride reduction of pH 6 to 8.

2. Suspend the washed support material containing aldehyde residues in an equal volume of 0.1-M sodium phosphate, pH 7.2, into which an amine-containing ligand has been dissolved. Notes: For protein ligands, a typical concentration range may be 3 to 5 mg/ml gel, but much higher concentrations may be used if a high density of coupled protein on the gel is desirable. In some cases, reactions containing protein at up to 20 mg/ml gel can be performed while still maintaining excellent coupling yields ( >85%). For instance, high-capacity protein A supports often are reacted at levels exceeding 10 mg protein A per ml gel to obtain maximal binding capacity for immunoglobulins. However, optimization of ligand density on the affinity support should always be done to avoid having too much or too little ligand present, which could cause significant nonspecific binding at the high end or unacceptably low capacity at the low end. High densities of immobilized protein could also result in an affinity support in which it is difficult to effect elution of the target molecule after binding. For small amine-containing affinity ligands, a concentration of 2 to 3 mg ligand/ml gel for the immobilization reaction may be sufficient to obtain good binding capacity on the resultant affinity gel. Alternatively, a concentration representing 5 to 10 times the concentration of reactive aldehydes on the matrix may be used to ensure a high density of the final coupled ligand. The ultimate concentration of ligand used in the reaction should be optimized by performing a series of coupling reactions at different initial ligand concentrations and determining the performance of the affinity supports in capturing and eluting the desired target molecule. The coupling of diamine spacer molecules should be done at high concentration (i.e., 1-M) to avoid internal crosslinking or bead aggregation during the coupling reaction.

3. Stir the gel/ligand slurry in a fume hood using a paddle stirrer or using a rotator to maintain constant mixing.

4. Add to the gel slurry 0.63 g of solid sodium cyanoborohydride (NaCNBH3; MW = 62.84) [toxic!] or 2 ml of 5-M NaCNBH3 in 1-N NaOH with stirring. If the alkaline solution of cyanoborohydride is used, check the pH and readjust to 7.2, if required. Continue the reaction for 4 h to overnight at room

temperature. The amount of cyanoborohydride added—whether as the pure solid compound or as the solution—results in a 50-mM solution in the final reaction slurry, which is in a total volume of 200 ml. A shorter coupling time may be sufficient for many ligands, but the optimal conditions to give acceptable yields for a particular ligand should be determined by doing a series of coupling reactions at different time points. Reactions at 4°C may also be done for thermally sensitive proteins or ligands, but reaction times likely will have to be extended to get the same level of coupling as a room temperature reaction.

5. Transfer the gel slurry to a sintered glass filter funnel (in the fume hood) and wash with several bed volumes of water to remove most of the unreacted ligand and reaction byproducts.

6. Unreacted aldehyde residues remaining on the support should be blocked by the addition of a small amine-containing compound, such as ethanolamine or Tris. Ethanolamine typically works best, and the compound is small enough to react with all remaining aldehydes to form hydrophilic terminal hydroxyl groups on the matrix upon coupling. Wash the support once with an equal volume of 1-M ethanolamine, pH 7.2, and then transfer the wet gel cake to a clean flask or vessel used for mixing the reaction. Add with stirring 100 ml of 1-M ethanolamine, pH 7.2, along with 0.63 g (or 2 ml of the 5-M NaCNBH3 solution) of sodium cyanoborohydride. Readjust the pH if necessary and continue the blocking reaction for 30 min at room temperature.

7. Wash the support extensively with water, 1-M NaCl, and again with water to remove all unreacted components from the gel. Additional wash solutions may be utilized to completely remove ligands that may have some nonspecific binding potential and remain noncovalently bound to the immobilized ligand, such as the use of acidic and alkaline washes as well as washes containing denaturants (such as guanidine). After washing, the affinity gel may be stored as a 50% aqueous slurry containing a preservative at 4°C.

(B) COUPLING AT pH 10

This protocol may be used to increase the amount of ligand coupled to the support using reductive amina-tion if the pH 7.2 protocol does not give sufficient cou-pling yields or does not result in high enough ligand density on the support. In this method, the formation of Schiff bases is first done at pH 10, where they occur more efficiently, and then the pH is adjusted to 7.2 for the reductive amination step. This two-step protocol typically results in much greater coupling yields than



the previous procedure, especially when immobilizing proteins.

1. Wash 100 ml of a periodate-oxidized support into coupling buffer (0.1-M sodium carbonate, pH 10) and drain to a wet cake. Other buffer components may be added to the coupling buffer, such as the addition of 0.15-M NaCl or alternative buffer salts; however, avoid amine-containing compounds like Tris, glycine, or imidazole, which will compete in the reaction.

2. Suspend the washed support material containing aldehyde residues in an equal volume of 0.1-M sodium carbonate, pH 10, into which an amine-containing protein or ligand has been dissolved. See the notes in step 2 of the previous pH 7.2 coupling protocol that contains suggestions as to the amount of ligand to add to the coupling reaction.

3. Stir the gel/ligand slurry for 4 h to overnight at room temperature using a paddle stirrer or using a rotator to maintain constant mixing. For sensitive proteins, the reaction may be carried out at 4°C, but may require additional time to form the Schiff bases.

4. Transfer the reaction slurry to a sintered glass filter funnel, drain the gel of excess solution, and wash with 2 × 100 ml of 0.1-M sodium phosphate, pH 7.2, to reduce the pH for the reduction reaction. Drain to a wet cake.

5. Transfer the gel cake back to a clean vessel and suspend it in 100 ml of 0.1-M sodium phosphate, pH 7.2, and stir using a paddle stirrer. Add to the gel slurry 0.63 g of solid sodium cyanoborohydride (NaCNBH3; MW = 62.84) [toxic!] or 2 ml of 5-M NaCNBH3 in 1-N NaOH. If the alkaline solution of cyanoborohydride is used, subsequently check the pH of the solution and readjust to 7.2, if required.

6. React with mixing at room temperature for 4 h or at 4°C overnight.


8. Aldehyde residues remaining on the support should be blocked by the addition of a small amine-containing compound, such as ethanolamine or Tris. Ethanolamine typically works best, and the compound is small enough to react with all remaining aldehydes to form hydrophilic terminal hydroxyl groups on the matrix upon coupling. Wash the support once with an equal volume of 1-M ethanolamine, pH 7.2, and then transfer the wet gel cake to a clean flask or vessel used for mixing the reaction. Add with stirring, 100 ml of 1-M ethanolamine, pH 7.2, along with 0.63 g (or 2 ml of the 5-M NaCNBH3 solution) of sodium

cyanoborohydride. Readjust the pH if necessary and continue the blocking reaction for 30 min at room temperature.

9. Wash the support extensively with water, 1-M NaCl, and again with water to remove all unreacted components from the gel. Additional wash solutions may be utilized to completely remove ligands that may have some nonspecific binding potential and remain noncovalently bound to the immobilized ligand, such as the use of acidic and alkaline washes as well as washes containing denaturants (such as guanidine). After washing, the affinity gel may be stored as a 50% aqueous slurry containing a preservative at 4°C.

Glutaraldehyde ActivationGlutaraldehyde is a 5-carbon, bis-aldehyde com-

pound that can be used to activate amine- or hydra-zide-containing supports to form reactive groups for the subsequent immobilization of amine-containing ligands. Given its structure, it may appear that the alde-hydes in this crosslinker might undergo reactions with two amines much the same as that seen using similar aldehyde-containing supports for immobilization using reductive amination (described previously). It also might be expected that a bifunctional glutaraldehyde molecule would form two Schiff bases with two amine-containing molecules, which could then be reduced with cyanoborohydride to form secondary amine link-ages. However, the nature of glutaraldehyde coupling is far more complex than a straightforward reduc-tive amination process using its terminal aldehydes. Glutaraldehyde undergoes significant transformation reactions in aqueous solution (and even in some sol-vents), resulting in a range of potential derivative forms (Figure 15.34), which can create a variety of intermedi-ary reactive species. These transitional forms can then have a number of potential reaction routes with amines beyond simple Schiff base formation. In this regard, glutaraldehyde activation of amine-containing chroma-tography supports and subsequent coupling to amine-containing ligands can potentially entail Schiff base formation with aldehydes, the Michael-type addition of amines to α,β-unsaturated sites within glutaraldehyde polymers, covalent reactions with a dimeric cyclic glu-taraldehyde species, or ring formation through an aldol condensation product which subsequently can react with amines to form cyclic quaternary pyridinium complexes (see Chapter 2, Section 4.4, and Chapter 5, Section 6.2) (Monsan, 1978; Migneault et al., 2004).

H

O

H

O

Glutaraldehyde



The formation of aldol condensation unsaturated constructs and the potential for creating cyclic hemi-acetal entities due to the intra- and intermolecular reactions of glutaraldehyde in aqueous alkaline solu-tion both may result in nondescript oligomerization and layering of the active intermediates within and on a support material during activation. Formation of these polymeric glutaraldehyde species, once started, is not easy to control and could cause clogging of pore structures within chromatography supports as well as difficulty in reproducing the activation and immobili-zation reactions. In most cases, however, through care-ful adjustment of the initial glutaraldehyde activation

conditions, side reactions such as polymerization can be limited and result in more defined reaction products that work well for immobilization of amine-containing affinity ligands.

In acidic or neutral pH conditions, glutaraldehyde can undergo an intramolecular cyclization reaction to form a hemiacetal ring (tetrahydro-2H-pyran-2,6-diol) that subsequently can polymerize through 6,6′-oxo link-ages to form what some people call it by the nonstan-dard name “polycycloglutaracetal”. The monomeric hemiacetal or the polymeric polycycloglutaracetal can react with amines by nucleophilic attack at the anomeric carbons bearing the hydroxyl groups (on both sides of a

FIGURE 15.34 The potential transformations of glutaraldehyde in aqueous solution are complex and create many different reactive species. Once water reacts with one of the aldehydes of glutaraldehyde, the molecule can cyclize to form a hemiacetal derivative, which can also polym-erize in solution to create polycycloglutaracetal. The acetal derivatives are reactive with amine-containing molecules through addition to the anomeric carbon atoms on each outer ring. Glutaraldehyde may also form a variety of unsaturated polymers at alkaline pH, which can undergo numerous transformations, including the creation of high-molecular-weight polymers that can precipitate out of solution. The cyclic acetal derivative can react with an amine-containing molecule by addition to the double bond on its left ring and through addition to the anomeric carbon atom on the right hemiacetal ring. Reactions with the unsaturated polymer form of glutaraldehyde typically occur through Michael-type addition to the double bonds along the polymer backbone.

H

O

H

O

+H2O

OHO OH

Cyclic hemiacetal

Glutaraldehyde

OHO OO OHn

Polymericcyclic hemiacetal

(polycycloglutaracetal)

H

O

O HO H

H

On

a,β-Unsaturated polymer

H

O

H

O

OH

O

H

O

OH

H

O

O HO H

H

O xOHy

RH

O

O H

H

OxOH

y

Carboxylate orCH2OH group

H

O

HO

HO

Cyclization

Cyclization

Cyclic acetal

+H2O

Polymerization

O

H

O

O

x

y

+

Addition products



single cyclic acetal ring or at both ends of the polymer). Subsequent bond formation results in a substitution reaction that creates a secondary amine linkage along with the loss of one molecule of water (Figure 15.35). The reactions of monomeric glutaraldehyde hemiac-etal may be one of the principal species responsible for stable ligand coupling without the need for reduction of Schiff bases, which normally would be necessary to stabilize aldehyde–amine complexes. This monomeric form may also be the primary type observed in immo-bilization reactions using an activation reaction with dilute glutaraldehyde done for a brief period of time (Betancor et al., 2006) (Figure 15.36). In this case, the initial activation step of an amine-containing chroma-tography support, such as monoaminoethyl-N-amino-ethyl (MANAE)–agarose, involves the reaction of one anomeric carbon of the cyclic acetal with the amine on the matrix to form a secondary amine bond. The other anomeric carbon atom of the acetal then can be used to couple amine-containing ligands after removal of excess glutaraldehyde species.

Under slightly more alkaline pH conditions (~pH 8.5), hemiacetal formation of glutaraldehyde can be

accompanied by aldol dimerization to result in a bicyclic derivative, which can undergo reactions at two sites within the molecule (Tashima et al., 1991) (Figure 15.37). Coupling of amine-containing mole-cules to this dimeric compound occurs by nucleophilic substitution at the anomeric hemiacetal carbon atom as well as through Michael-type addition to the double bond on the aldol addition ring product (Figure 15.38). This dimeric form of glutaraldehyde may be the pre-dominant species observed in some activation and immobilization reaction protocols, wherein the activa-tion step is done by the addition of a relatively high concentration of glutaraldehyde that is reacted for an extended period of time (Betancor et al., 2006). This glutaraldehyde dimer is an extremely potent reactant with amines and it has been implicated as the form responsible for the best crosslinking performance in tissue fixation, protein conjugation, and in the immo-bilization of enzymes (Robertson and Schultz, 1970; Makino et al., 1988).

Dimeric cyclic glutaraldehyde can be used to acti-vate amine-containing chromatography supports, such as a MANAE–agarose matrix, to link initially to

OHO OH

Cyclic hemiacetal

OHO OO OHn

Polymericcyclic hemiacetal

(polycycloglutaracetal)

H2N

ONH

OH

NH2

H2N

ONH

NH

ONH

OO OHn

NH2

ONH

OO NHn

Amine-containingmolecule


Amine-containingmolecul


FIGURE 15.35 The glutaraldehyde cyclic hemiacetal can react with two amine-containing molecules to result in addition to the anomeric carbon atoms on both sides of its ring structure.



the α,β–unsaturated double bond to create an inter-mediate reactive derivative. The initial activation reaction is done at relatively high concentrations of glu-taraldehyde at neutral pH and for an extended period to ensure that the reaction occurs with a glutaraldehyde dimer instead of a hemiacetal monomer (Betancor et al., 2006). The activated support then can be coupled with amine-containing ligands through covalent attachment to the anomeric carbon of the hemiacetal ring with loss of one molecule of water to form a secondary amine linkage (Figure 15.39). Both of these reactions require no

reducing step as is common with coupling aldehydes to amine-containing molecules, and the coupling occurs rapidly and in high yield.

In even more alkaline pH conditions, glutaralde-hyde can undergo transformation by a series of aldol condensation reactions that creates, after dehydration and loss of the hydroxyl groups along with the adja-cent hydrogens, numerous α,β-unsaturated sites within glutaraldehyde polymers of indeterminate length (Rembaum et al., 1978). The resultant polymers report-edly also can contain a number of carboxylic, hydrox-ylic, and aldehyde groups along its length, although the exact composition is not well characterized (Margel and Rembaum, 1980). The presence of carboxylate func-tionalities presumably is due to further redox reactions that occur with the initial polymer in the presence of oxygen. Aldol polymerization of glutaraldehyde at pH 11 or above reportedly can be so severe that precipita-tion occurs in aqueous solution (Monsan et al., 1975). Kawahara et al. (1992) indicated that extensive polymer-ization also can occur if the glutaraldehyde is dissolved in anhydrous solvent as well as high pH aqueous solu-tions. This form of polyglutaraldehyde long has been used to activate amine-containing microparticles for the subsequent coupling of amine-containing ligands (Rembaum et al., 1978; Margel and Rembaum, 1980) (see also Chapter 14). However, it is likely that the other

OHO OH

Glutaraldehydecyclic hemiacetal

O

HN

NH2

MANAE-modifiedsupport containing

primary amines

+

O

HN

ONH

OH

Glutaraldehyde-activatedintermediate

O

HN

ONH

H2N

NH

Amine-containingligand

Ligand coupling viasecondary amine bond

FIGURE 15.36 The activation of MANAE–agarose with glutaral-dehyde monomer (under dilute glutaraldehyde conditions) and cou-pling of an amine-containing ligand.

OH OOH O

OO OHO

O

O

O

O

O OH

O

OH O

O

Glutaraldehydemolecules

Dimerization Cyclization

Bicyclization Elimination

FIGURE 15.37 Formation of a dimeric, bicyclic glutaraldehyde.

O OH

O NH2 H2N

O OH

O HN

O NH

O HN

Dimericglutaraldehyde


Activatedintermediate


Conjugateformation

FIGURE 15.38 Reaction of glutar-aldehyde dimer with two amine-con-taining compounds.



monomer and dimer species described above may also contribute to the overall coupling reactions occurring with glutaraldehyde, depending on the exact activation procedure being used.

Aldol-mediated polyglutaraldehyde formation can be used to activate amine-containing supports by ini-tially coupling through Michael-type addition reactions to various double bonds within the polymer strands. After activation with an excess of this polymeric form, the remaining uncoupled polyglutaraldehyde can be washed off the matrix and the activated intermedi-ate used to immobilize an amine-containing ligand (Figure 15.40). Since polyglutaraldehyde contains both aldehydes and double bonds, activation of supports with this species likely involves two routes of amine reactivity. The Michael-type addition of amines to the double bond sites will result in secondary amine link-ages, which have no requirement for further stabiliza-tion by reduction. However, amines may also link to the polymer through Schiff base formation with the aldehydes still present in each polymer strand. These Schiff bases may need a reductive amination reac-tion using sodium cyanoborohydride to stabilize them against hydrolysis and form secondary amine bonds. Particularly, if ligands are being immobilized that con-tain only a single amine group, then it is recommended that reduction be carried out to prevent ligand leakage through hydrolysis.

Glutaraldehyde and its derivatives that form in aque-ous solution potentially can react with a number of nucleophilic groups on proteins or other ligands, such as amines, thiols, and imidazole nitrogens, as well as phenolic –OH groups (Habeeb and Hiramoto, 1968). The relative reactivity of glutaraldehyde toward some of the most predominant functional groups in proteins has been estimated to be ε-amine > α-amine > guanidi-nyl > secondary amine > hydroxyl groups (Migneault et al., 2004). Thiol reactivity with glutaraldehyde-reactive species is also possible, but it has been reported to occur only in the presence of amine functionalities (Okuda et al., 1991). The bifunctional nature of the glutaralde-hyde monomeric or dimeric forms makes it suitable for activation of chromatography supports followed by immobilization of affinity ligands containing nucleo-philes, such as amines. Due to the diverse reactions of glutaraldehyde in aqueous solution, the stability of coupling to amine-containing ligands typically is far greater than the stability of the initial Schiff bases cre-ated from aldehyde-containing supports as they react with amine-containing ligands (see previous section). Normally without reductive amination coupling of amino ligands to aldehyde-containing supports, the immobilized ligand will leach off the matrix as the Schiff bases slowly hydrolyze in aqueous solution. By contrast, glutaraldehyde immobilization is more resistant to ligand leakage even without a subsequent

O OH

O

O

HN

NH2


primary amines

+

Dimeric glutaraldehyde

O

OH

O

O

HN

NH

Glutaraldehyde-activated support

H2NAmine-containingligand

O

NH

O

O

HN

NH

Immobilized ligandthrough secondary

amine linkage

FIGURE 15.39 Activation of MANAE–agarose with dimeric glutaraldehyde and coupling of an amine-containing ligand.



reduction step, because most of the ligand coupling reactions involve addition reactions without the inter-mediate formation of a Schiff base.

Due to the diverse nature of glutaraldehyde reac-tions, the exact reactive species that is formed after activation may be less certain than when using other activation methods. Nevertheless, the immobiliza-tion reaction can be designed to take place with high efficiency and yield. In addition, recently it has been shown that by carefully controlling the level of glutar-aldehyde activation of an amine-containing support at neutral pH, the result can be fairly well constrained to either a highly reactive dimeric species or a somewhat less reactive and predominantly monomeric glutaralde-hyde derivative (Betancor et al., 2006). The major reac-tive species created on an amine-containing support can be predetermined by the amount and concentration of glutaraldehyde addition as well as by the time of the reaction. By contrast, activation of an amine-containing support under highly alkaline conditions will result in

a greatly polymerized glutaraldehyde intermediate that reacts with nucleophiles on ligands mainly through the addition to double bonds (although Schiff base forma-tion may also occur).

The use of glutaraldehyde to activate and couple enzymes for the preparation of immobilized reactors remains one of the most popular methods for prepara-tion of these catalytic supports (Burteau et al., 1989; Van Aken et al., 2000; Dos Reis-Costa et al., 2003; Magnan et al., 2004; Seyhan and Alptekin, 2004; Betancor et al., 2006; Ma et al., 2011; Matosevic et al., 2011). Unlike reductive amination processes described in the previ-ous section, immobilization using glutaraldehyde usu-ally does not require the use of a reducing agent, such as the hazardous sodium cyanoborohydride. Especially when coupling multivalent protein molecules, immo-bilization onto a glutaraldehyde-activated support can create multiple attachment sites that firmly link the ligand with minimal potential for leakage. Thus, if care is taken to control the initial activation reaction,

H

O

O HO H

H

On

O

HN

NH2


primary amines

+O

HN

NH

Glutaraldehyde-activated support

H2NAmine-containingligand

H

O

O

H

O

H

HO

n

O

HN

NH

H

O H

H

HO

nN

N

Polyglutaraldehyde

Ligand couplingvia Schiff base

formation

FIGURE 15.40 Activation of MANAE–agarose with polyglutaraldehyde (aldol-mediated) followed by coupling of an amine-containing ligand to form a secondary amine linkage.



the resultant preparation of an affinity support can be performed reproducibly and give high activity in its intended application.

The following protocols can be used to activate amine-, amide-, or hydrazide-containing supports with glutaraldehyde for subsequent coupling of amine-con-taining ligands. The first two procedures are based on the method of Betancor et al. (2006). These activations can be performed using any chromatographic support mate-rial containing available primary amines, such as the use of an agarose support that has been modified with a diamine spacer to contain free terminal amino groups. One such support described in the literature is mono-aminoethyl-N-aminoethyl (MANAE)–agarose, which contains low-pKa amino groups that are highly reac-tive using neutral pH buffers (Fernandez-Lafuente et al., 1993). In one regard, the final level of glutaraldehyde activation can be controlled by regulating the amount of these amines produced initially on the support material.

Glutaraldehyde should be handled with care in a fume hood while using the appropriate personal pro-tective equipment to prevent contact or exposure to the solutions or vapors. All waste should be treated as aqueous hazardous waste and disposed of according to environmental regulations and local policy.

ACTIVATION PROTOCOLS

(A) Glutaraldehyde Activation to Form a Monomeric-Reactive Species

This protocol forms a reactive intermediate with mod-erate rates of reactivity for coupling amine-containing ligands.

1. Wash 100 ml of an amine-containing support material using gentle vacuum filtration in a sintered glass filter funnel (such as MANAE–agarose, which can be prepared by methods described in this chapter for coupling diamine spacers to chromatography supports) with several bed volumes of deionized water to remove any storage solution. Finally, wash the support with 200 ml of activation buffer (0.2-M sodium phosphate, pH 7.0). The support material and all wash solutions as well as the reaction should be maintained at 25°C. Note: Using a buffer pH of >8 will result in glutaraldehyde polymerization and an uncontrolled reaction.

2. In a fume hood, add the moist gel cake to a clean vessel that can be used to stir or rotate the gel during the activation and coupling reactions. Add 200 ml of 0.5% (v/v) glutaraldehyde dissolved in coupling buffer with mixing to resuspend the support in the solution.

3. React with mixing for 1 h.4. Remove the gel slurry from the reaction vessel

and drain the excess glutaraldehyde solution into

a suction filter flask using vacuum filtration in a sintered glass filter funnel (in a fume hood). Wash the activated support extensively with 25-mM sodium phosphate, pH 7.0, buffer and then with deionized water. The activated support can be stored at 4°C until ready for ligand coupling.

(B) Glutaraldehyde Activation to Form a Dimeric-Reactive Species

This protocol forms a reactive intermediate with extremely high reactivity for coupling amine-containing ligands.

1. Wash 100 ml of an amine-containing support material using gentle vacuum filtration in a sintered glass filter funnel (such as MANAE–agarose) with several bed volumes of deionized water to remove any storage solution. Finally, wash the support with 200 ml of activation buffer (0.2-M sodium phosphate, pH 7.0). The support material and all wash solutions as well as the reaction should be maintained at 25°C.

2. In a fume hood, add the moist gel cake to a clean vessel that can be used to stir or rotate the gel during the activation and coupling reactions. Add 200 ml of 15% (v/v) glutaraldehyde dissolved in coupling buffer with mixing to resuspend the support in the solution.

3. React with mixing for 15 h at 25°C.4. Remove the gel slurry from the reaction vessel

and drain the excess glutaraldehyde solution into a suction filter flask using vacuum filtration in a sintered glass filter funnel (in a fume hood). Wash the activated support extensively with 25-mM sodium phosphate, pH 7.0, buffer and then with deionized water. The activated support can be stored at 4°C until ready for ligand coupling.

IMMOBILIZATION OF LIGANDS ON GLUTARALDEHYDE-ACTIVATED SUPPORTS

The coupling of amine-containing ligands such as pro-teins or enzyme reactors to glutaraldehyde supports pre-pared according to the previous activation protocols will proceed rapidly under low ionic strength conditions (i.e., using 25-mM potassium phosphate, pH 7, with no addi-tional salt added). Under high ionic strength conditions (25-mM potassium phosphate, pH 7, containing 0.5-M NaCl) the dimeric glutaraldehyde activation process (protocol (B), above) will react and couple ligand almost as fast as when using low ionic strength buffers; how-ever, when using the monomeric glutaraldehyde acti-vation process (protocol (A), above) it results in slower coupling rates, which could take many hours or even overnight to go to completion (Betancor et al., 2006). The reason coupling takes place faster under low salt condi-tions is thought to be due to the formation of an initial ionic interaction between the protein molecules to be



coupled and the underlying secondary amines to which the glutaraldehyde derivatives first are linked during activation. The initial ion-exchange character of the acti-vated support facilitates recruitment of the protein to the immediate proximity of the reactive groups, thus accel-erating immobilization. Adding high salt obviates these charge interactions and thus slows down the overall reaction process.

Protocol1. Wash 100 ml of a glutaraldehyde-activated support

containing dimeric glutaraldehyde (activation protocol (B), above) with 2 bed volumes of 25-mM potassium phosphate, pH 7.0 (coupling buffer), and drain to a wet cake. The addition of salt (NaCl) to the coupling buffer may be done for ligands that require it, but the use of additional ionic strength may also require increased reaction times to accommodate a somewhat slower reaction rate (see previous discussion). Transfer the support to a vessel that can be used to mix the gel suspension during the coupling reaction by end-over-end rotation or by using a paddle stirrer.

2. Dissolve an amine-containing ligand such as a protein, enzyme, or small ligand in 100 ml of coupling buffer. The ligand should be at a concentration of at least 2-fold greater than the quantity desired to be immobilized. For protein ligands, reacting at a concentration of 3 to 5 mg/ml resin is often sufficient. However, if a high density of immobilized protein or enzyme is desired, reactions can be done at concentrations of 10 to 20 mg/ml to obtain better coupling yields. For small ligands, the reaction should be done at a concentration that exceeds the level of glutaraldehyde activation or at least by using a 2-fold greater amount of ligand than the anticipated final immobilization level. To obtain an optimal affinity support for a given application, it may be necessary to perform a series of reactions using different ligand concentrations to determine the best coupling concentration to use.

3. Mix the ligand solution with the activated support and mix to resuspend the resin.

4. React with mixing for 2 h at room temperature or at 25°C. If a monomeric glutaraldehyde-activated support is used (activation protocol (A), above), then the reaction time should be extended by at least an hour. If high salt was added to the coupling buffer (e.g., 0.5-M NaCl) in step 1 and 2, then the reaction using dimeric glutaraldehyde activation should be extended to 3 h, and if monomeric glutaraldehyde activation was used with high salt, it should be extended to an overnight reaction.

5. Remove the reaction slurry and transfer to a filter flask. Drain the excess reaction solution

and wash with water to remove the majority of uncoupled ligand. If the remaining active groups are not to be blocked, then the wash should be done extensively with water, 1-M NaCl, and again with water.

6. Reactive groups remaining on the support should be blocked by the addition of a small amine-containing compound, such as ethanolamine or Tris. Ethanolamine typically works best, and the compound is small enough to react with all remaining active sites to form hydrophilic terminal hydroxyl groups on the matrix upon coupling. In a fume hood, wash the support once with an equal volume of 1-M ethanolamine, pH 7.0, and then transfer the wet gel cake to a clean flask or vessel used for mixing the reaction. Add with stirring, 100 ml of 1-M ethanolamine, pH 7.0, along with 0.63 g of sodium cyanoborohydride (or 2 ml of a 5-M NaCNBH3 solution in 1-N NaOH). Cyanoborohydride is extremely toxic and should be handled with care in a fume hood. The use of a reducing agent in the final blocking step is optional, but it is included here to ensure that if there are any aldehydes present in the activated support they too will be blocked by the ethanolamine addition (by reducing any Schiff bases formed by the reaction of the amine on ethanolamine with the aldehyde groups). Note that if the ligand is sensitive to the presence of cyanoborohydride, use the ethanolamine solution without adding the reducing agent. Readjust the pH if necessary and continue the blocking reaction for 30 min at room temperature.

7. Wash the support extensively with water, 1-M NaCl, and again with water to remove all unreacted components from the gel. Additional wash solutions may be utilized to completely remove ligands that may have some nonspecific binding potential and remain noncovalently bound to the immobilized ligand, such as the use of alternating acidic and alkaline washes as well as washes containing denaturants (such as guanidine). After washing, the affinity gel may be stored as a 50% aqueous slurry containing a preservative at 4°C.

NHS Ester and NHS Carbonate ActivationN-Hydroxysuccinimide (NHS) esters are activated

carbonyl groups that may be formed from carboxyl-ates through the use of a carbodiimide-mediated esteri-fication reaction done under aqueous or nonaqueous conditions. They may also be formed through trans-esterification or transfer reactions using reagents such as Sakakibara’s reagent, which is the NHS ester of triflu-oroacetate, TSTU (N,N,N’,N’-tetramethyl(succinimido)uronium tetrafluoroborate), or disuccinimidyl carbonate



(DSC) (Sakakibara and Inukai, 1965; Wilchek et al., 1994). NHS esters are electrophilic reactive groups that can couple with nucleophiles such as amines to form stable amide bonds. They have been used extensively in the design of crosslinkers and modification reagents to form covalent conjugates with amine-containing pro-teins or other amine-containing molecules (Chapter 3, Section 1.4). NHS esters also can be used to immobi-lize amino-ligands onto chromatography supports in a rapid, single-step reaction that typically occurs in high yield (Prokudina et al., 2011) (Figure 15.41).

Chromatography supports containing NHS esters are available commercially from several sources, and most contain a spacer arm that terminates in the activated car-boxylate (e.g., Thermo Fisher, Bio-Rad, GE Healthcare). These supports thus are synthesized first by building a spacer arm off the chromatography support and then forming the NHS esters by activation of the carboxyl-ates. A wide range of spacer molecules can be considered for this purpose, including hydrophobic aliphatic com-pounds such as 6-aminocaproic acid or hydrophilic spac-ers such as those containing PEG-based cross-bridges (i.e., amino–PEG4–carboxylate; Thermo Fisher, Quanta BioDesign). Usually, the spacers are attached to the sup-port through their amino end by coupling to an amine-reactive resin material, such as any of the amine-reactive immobilization chemistries described in this chapter. Once the carboxylate spacer derivative is formed on the support, it then can be activated to create the NHS ester groups using a number of different reaction routes.

When working with microparticles or nanoparticles, the formation of the NHS ester reactive group often is done in aqueous solution immediately prior to add-ing an amine-containing molecule to the particles for coupling (Chapter 14). This is accomplished by using a water soluble carbodiimide such as EDC (Chapter 4, Section 1) in the presence of NHS or sulfo-NHS (the sulfonated form promotes greater water solubility). The active NHS esters are formed despite the fact that the aqueous environment also causes continual hydro-lysis of the esters back to unreactive carboxylates. This activation strategy works well with small nonpo-rous particles if the ligand to be immobilized is added immediately after the activation step or even during the

activation process. However, when working with chro-matography supports that are made of larger porous particles, it is best to create the NHS ester groups in a nonaqueous environment so that competing hydro-lysis is eliminated and maximal activation levels are obtained throughout the inner bead structures.

Although the use of a carbodiimide-mediated for-mation of NHS ester groups is common when work-ing with carboxylate microparticles or nanoparticles, a major potential side reaction has been reported in the formation of NHS esters using carbodiimides on hydroxylic supports that have been modified to pos-sess carboxylate spacers (Wilchek and Miron, 1987). If a reaction involving the carbodiimide DCC with added NHS is used to create the NHS ester groups on a car-boxylate support under nonaqueous conditions, as is also typical for synthesizing most NHS ester-containing bioconjugation reagents, the formation of a bis-NHS derivative of β-alanine can be created at the same time that NHS esters are being formed with the carboxyl-ate spacers on the matrix. This side reaction results from the initial interaction of DCC with the hydroxyl on NHS to form a carbodiimide-activated NHS group. This ability of carbodiimides to react with hydroxyl groups correlates to the reported activation of hydrox-yls on chitosan using the carbodiimide EDC (Chiou and Wu, 2004). The DCC–NHS intermediate in this side reaction goes on to react with two additional equiva-lents of NHS, which then is followed by a Lossen rear-rangement to create a reactive carbamate ester on the amino end of β-alanine and an NHS ester on the car-boxylate end (Gross and Bilk, 1968) (Figure 15.42). This bifunctional compound then reacts with any remain-ing hydroxyl groups within the original matrix to cre-ate unstable ester linkages. The other end still can react with amine-containing ligands to form a covalent bond, but the immobilized product formed from this side reaction is unstable due to the potential for ester hydro-lysis, and thus it can continually leach off the matrix in aqueous solution (Figure 15.43). This side reaction may be the main reason why many NHS ester-mediated immobilization reactions result in leaky supports.

To overcome these potential side reactions a two-step protocol might be used wherein the carboxylate

Support containingNHS ester-reactive groups

O

ON

O

O

+ H2N

Amine-containing

ligand

O

HN

Immobilization ofligand through

amide bond formation

FIGURE 15.41 Reaction of an NHS ester-activated support with amine-containing ligands to form an amide bond.



support first is activated with a carbodiimide such as DCC in nonaqueous conditions, then the support is washed free of excess carbodiimide and reacted with NHS to form the final NHS esters. However, this pro-cedure has been found to result in decreased activation yields and is cumbersome (Wilchek and Miron, 1987). Fortunately, there are now alternative strategies to cre-ating NHS esters on hydroxylic support materials that have the ability to give highly stable reaction prod-ucts without the potential side reactions that carbodi-imides plus NHS are known to cause. One route was developed by Wilchek et al. (1994) specifically to avoid the problems of using carbodiimides with NHS and hydroxylic supports.

The coupling reagent TSTU originally was devel-oped as an amide bond-forming agent for peptide synthesis (Knorr et al., 1989; Bannwarth and Knorr, 1991). It is able to directly activate a carboxylate group

to an NHS ester in a single step without the forma-tion of detrimental side reaction products or other reactive components having crosslinking potential. Uronium coupling reagents react with carboxylate groups through an initial attack on the central posi-tively charged carbon atom of the tetramethyl uronium component (Figure 15.44). This is followed by a rear-rangement leading to the formation of an intermediary uronium ester with the carboxylate along with displace-ment of the leaving group attached to the uronium part of the reagent. In the case of TSTU, it is the NHS group that gets removed in this step. Next, the hydroxyl of the released NHS group in turn attacks the carbonyl car-bon of this intermediate uronium ester, which causes displacement of the tetramethyl uronium group and formation of the final NHS ester on the carboxylate. Most TSTU activation reactions are done in nonaque-ous solvent (e.g., dioxane or DMF), which preserves the

N

O

O

OH

NHS

+

NCN

R

R

Carbodiimidecompound

N

O

O

ONH

N

R

R

Activated esterintermediate

N

O

O

ONH

N

R

R

N

O

O

OH

NH2

O

O

HONH

N

R

R

ON

O

O

N

O

ON

O

O

N

O

O

HO

CO

O NH

O

O

ONN

O

O O

O

Attack of secondNHS molecule

Attack of thirdNHS molecule

Ring cleavageand rearrangement

N-(Succinimidooxy)carbonyl]-beta-alanineN-hydroxysuccinimide ester, constructed

from three NHS molecules

FIGURE 15.42 Reaction of NHS with DCC to form an activated β-alanine byproduct, which can form when using carbodiimides to create NHS esters in organic solvent.



NHS ester from hydrolysis; however, the reagent may also be used in a mixture of organic solvent and water. In addition to Bannwarth and Knorr (1991) demonstrat-ing an organic/water reaction environment consisting of a 2 :2 :1 mixture of DMF/dioxane/water, Andersson et al. (1993) used the coupling reagent to activate a carboxyl-ate derivative of a saccharide in an aqueous/organic sol-vent mixture and used the NHS ester to conjugate the saccharide to a protein.

N O+ N

NO

O

B–FF

F

F

TSTU2-succinimido-1,1,3,3-tetramethyl-

uronium tetrafluoroborate(tetrafluoroborate salt)

ONH

O

O

ON N

O

OO

O

O–

O

NH

OH

Support containingcarboxylate-terminalspacers and excess

hydroxyls O

O

NH

OH

Carbodiimide

+

bis-NHS–beta-alanine

N

O

ONHS ester-activated

carboxylate

O

O

NH

N

O

O

NHS(nonaqueous)

OHN

OO

O

N

O

OUnstableester bond

FIGURE 15.43 Reaction of activated β-alanine with the hydroxyl groups remaining on a support, which was modified to contain carboxylates.

N + +

+

ON

NO

O

O

O–

Suppport containingcarboxylate groups

TSTU

+ O

NO

N

OHN

O

O

+ O

O N

O

O

N OH

N

NHS ester-activated support

Intermediate uroniumester formation followed

by attack of NHS

FIGURE 15.44 Mechanism of NHS ester formation using TSTU.



In addition to the use of TSTU to create NHS ester-activated derivatives, the bifunctional reagent N,N’-disuccinimidyl carbonate (DSC) also can be used to form these reactive groups on carboxylate-containing matrices (Ogura et al., 1979; MacBeath and Schreiber, 2000). In this reaction, which is done in a nonaqueous environment, DSC reacts directly with the carboxylate to transfer one of its NHS groups to the acid in a transesterification process (Figure 15.45). The result is the single-step for-mation of an NHS ester on the carboxylate-containing support, which is then washed free of excess reactants and stored in nonaqueous solution to maintain stability of the ester until use. The byproducts of this process are CO2 and NHS, both of which are completely innocuous. This reaction also avoids the potential side reactions of a carbodiimide/NHS reaction to form NHS esters. In simi-lar methods, other bis-carbonates have been used to acti-vate carboxylates, such as tert-butyl carbonates (Basel and Hassner, 2002). The formation of NHS esters on car-boxylate supports with bis-carbonates can be performed in anhydrous DMF (or other appropriate water-miscible solvents) using 100-mM DSC along with an equal con-centration of an organic base as a proton acceptor, such as N,N-diisopropylethylamine (DIEA).

As an alternative approach, reactive NHS esters also can be created directly on hydroxyl-containing chroma-tography supports without the need to add an interme-diary carboxylate spacer. In this strategy, the hydroxylic resin is washed into nonaqueous solvent and activated using the reagent DSC. This compound reacts with a hydroxyl on the matrix with loss of one NHS group to form an N-succinimidyl carbonate reactive group, which is similar in reactivity toward amine nucleophiles as that of an NHS ester formed from a carboxylate (Wilchek and Miron, 1985). This same activation pro-cess has been used successfully to activate the hydroxyl groups on PEG compounds for subsequent coupling

to amine-containing molecules (Chapter 18, Section 2.1). The immobilization of amine-containing ligands onto DSC-activated supports results in the formation of carbamate linkages (Figure 15.46), which are identi-cal to the bonds formed from the reaction of amines with CDI-activated matrices (see this chapter, CDI-Activated Supports). Carbamates have been found to be very secure linkages that are similar to amide bonds in chemical stability. Another advantage of DSC activation of hydroxyl supports is that the hydroxyl is reformed by hydrolysis if an amine ligand is not coupled during the immobilization reaction. This leaves an uncharged, hydrophilic group on the matrix if hydrolysis occurs as opposed to a typical NHS ester, which upon hydrolysis, creates a carboxylate group bearing a negative charge. Thus, if a spacer arm on the matrix is not required to present the affinity ligand to its intended binding target, then direct activation of a hydroxylic support with DSC potentially is a better choice than an NHS ester formed from a carboxylate spacer. Even if a spacer arm is desir-able to create an immobilized ligand, the use of a PEG spacer containing a terminal hydroxyl group could be created to subsequently activate the spacer with DSC. The use of PEG spacers would form a highly hydro-philic environment that will dramatically lower the nonspecific binding potential of the final affinity sup-port. In addition, if a hydroxylic matrix first is modi-fied to contain carboxylate spacers for conversion to NHS esters using DSC, it is likely that some remain-ing hydroxyls also are converted to the NHS carbonate in the activation process. Therefore, this result would produce both amide and carbamate bonds in the final immobilized ligand support, the relative proportion of which would depend on the ratio of activated carboxyl-ates to activated hydroxyls.

A final option using DSC for the activation of chroma-tography supports is to use it to activate amine-containing

O

OH


+

O

O

ONN

O

O O

O

Disuccinimidylcarbonate (DSC)

O

O N

O

O

+HO

N

O

O

+ CO2

NHS ester-activated support

NHS

Nonaqueous

FIGURE 15.45 DSC activation of carboxylate supports to form NHS esters.



matrices. A beaded chromatography support modi-fied to contain amine spacers can be created through the coupling of a diamine compound to an activated matrix, which then results in terminal primary amines on the support for further activation and coupling reac-tions (see section on coupling spacer molecules). DSC can be used to directly activate these amines to reac-tive NHS carbamates, which in turn can be coupled to amine-containing ligands to form an isourea link-age (a urethane bond) (Figure 15.47). This reaction has been used successfully to couple immobilized proteins with other amine-containing molecules (MacBeath and Schreiber, 2000) and to activate amine spacers on a slide surface for immobilization of amine-containing ligands (Niemeyer, 2004). This procedure offers a way of coupling amine-containing ligands to an amine-containing support without further modification of the amino spacers to convert them to a different functional-ity for activation. However, it should be kept in mind that the underlying amines on this support may create positively charged sites, especially with regard to any DSC-activated amines that do not couple to ligands in the immobilization reaction, because the hydrolysis product regenerates the original amine. Thus, using this method there may be some potential for nonspecificity

in the final affinity support from ion-exchange effects due to leftover or uncoupled amines.

The following protocols represent methods that can be used to create amine-reactive NHS esters on chroma-tography supports. Carbodiimide-based methods using DCC in organic solvent can also be done, but are not included in the recommended protocols due to the likely side reactions involving the formation of the bifunctional β-alanine crosslinking agent described previously, which would result in an affinity support having a high poten-tial for ligand leakage. Note that for protocols related to the activation and coupling of ligands to carboxylate-containing microparticles or carboxylated nanoparticles using the carbodiimide EDC with NHS to form interme-diate NHS esters, the reader should refer to Chapter 14.

All solvents used in the following protocols should be as anhydrous as possible to avoid the potential for hydrolysis of the NHS ester groups during storage of the activated supports.

ACTIVATION PROTOCOLS

(A) NHS Ester Activation Procedure using TSTUThe following procedure is based on the method

of Wilchek et al. (1994) using TSTU in a reaction with

OH

Suppport containinghydroxyl groups

+

O

O

ONN

O

O O

O


O+

HON

O

O

NHS carbonate-activated support

NHS

Nonaqueous

O

ON

O

O

H2N


O+

HON

O

O

Ligand immobilizedvia carbamate bond

NHS

O

HN

Aque

ous

orno

naqu

eous

FIGURE 15.46 DSC activation of hydroxyl supports followed by coupling to amine-containing ligands to form carbamate bonds.



carboxylate groups on a chromatography support to form reactive NHS esters that can be used to couple with amine-containing ligands.

1. Wash 100 ml of a hydroxylic chromatography support material (such as crosslinked agarose) that has been modified to contain carboxylate groups with 0.3-N HCl and then with water (several bed volumes each) using a sintered glass filter funnel. In a fume hood, sequentially wash the support into increasing concentrations of dioxane-in-water (such as 25% (v/v), 50%, and 75% dioxane/water) and finally with 100% solvent for at least 10 to 20 bed volumes to remove the last traces of water. Drain to a wet gel cake and then pull a gentle vacuum on the filter funnel while breaking up the gel cake with a spatula to obtain moist, evenly divided pieces, which still contain dioxane within the internal pores of the beads. Remove the vacuum as soon as the gel is uniformly divided and appears as a white fluffy snow-like consistency. Be careful not to allow the support to dry, as this will cause particle collapse and irreversible damage to the pore structure.

2. In a fume hood, prepare a 100-ml solution of 0.1-M TSTU (N,N,N’,N’-tetramethyl(succinimido)uronium

tetrafluoroborate) in DMF. Add the washed gel cake to the TSTU solution with stirring to resuspend the gel. Next add 0.244 g of 4-(dimethylamino)pyridine (DMAP) to the gel suspension to prepare a 0.2-M DMAP solution and mix to dissolve. A small amount of additional DMF may be added if necessary to aid in stirring the mixture.

3. Mix the reaction for 1 h by rotation or using an overhead paddle stirrer.

4. Wash the activated support with several bed volumes each of DMF, methanol, and isopropanol to remove excess reactants and reaction byproducts. The NHS ester-activated support can be stored as a 50% slurry in isopropanol at 4°C until use.

(B) NHS Ester Activation of Carboxylate Supports using DSC1. Wash 100 ml of a carboxylate-containing

chromatography support material (such as crosslinked agarose that has been modified to contain carboxylate groups) with water (at least several bed volumes) using a sintered glass filter funnel. In a fume hood, sequentially wash the support into increasing concentrations of

NH2

Suppport containingamine groups

+

O

O

ONN

O

O O

O


HN

+HO

N

O

O

NHS carbamate-activated support

NHS

Nonaqueous

O

ON

O

O

H2N


HN

+HO

N

O

O

Ligand immobilizedvia isourea linkage

NHS

O

HN

Aque

ous

orno

naqu

eous

FIGURE 15.47 Activation of amine-containing supports with DSC and subsequent coupling of amine-containing ligands.



acetone-in-water (such as 25% (v/v), 50%, and 75% acetone/water) and finally with 100% dry acetone for at least 10 to 20 bed volumes to remove the last traces of water. Alternative solvents include dioxane, DMF, DMAC, or DMSO, which can be substituted for acetone depending on the compatibility of the matrix to a particular solvent. Drain to a wet gel cake and then pull a gentle vacuum on the filter funnel while breaking up the gel with a spatula to obtain moist, evenly divided pieces, which still contains acetone within the internal pores of the beads. Remove the vacuum as soon as the gel is uniformly divided and appears as a white fluffy snow-like consistency. Be careful not to allow the support to dry too much, as this will cause particle collapse and irreversible damage to the pore structure.

2. In the fume hood, prepare 100 ml of a solution consisting of 80 mg/ml DSC in dry acetone. Stir vigorously to dissolve. There may be some insoluble DSC left in the solution as a fine suspension, but this material will be driven into solution as the activation reaction proceeds.

3. Add the acetone wet gel cake to the DSC solution with stirring to uniformly resuspend the support. To this suspension, add 6.5 g of DMAP to provide a base to catalyze the reaction; alternatively, add 7.5 ml of anhydrous triethylamine (TEA). Mix the gel suspension using an overhead stirring paddle or by end-over-end rocking in the fume hood.

4. React for 1 h at room temperature with mixing.5. In the fume hood, wash the activated support with

acetone to thoroughly remove excess reactant and reaction byproducts. Wash with at least 10 bed volumes of solvent to ensure removal of the last traces of unreacted DSC.

6. Store the NHS ester-activated support as a 50% slurry in dry acetone at 4°C until use.

(C) NHS Carbonate Activation of Hydroxylic Supports using DSC1. Wash 100 ml of a hydroxyl-containing

chromatography support material (such as agarose, Toyopearl, or Trisacryl) with water (3–5 bed volumes) using a sintered glass filter funnel. In a fume hood, sequentially wash the support in increasing concentrations of acetone-in-water (such as 25% (v/v), 50%, and 75% acetone/water) and finally with 100% dry acetone for at least 10 to 20 bed volumes to remove the last traces of water. Alternative solvents include dioxane, DMF, DMAC, or DMSO, which can be substituted for acetone depending on the compatibility of the matrix to a solvent. Drain to a wet gel cake and then pull a gentle vacuum on the filter funnel while breaking up the gel cake with a spatula to obtain moist, evenly divided pieces, which

still contain acetone within the internal pores of the beads. Internal pores of the beads, with a spatula to obtain moist, evenly divided pieces. Remove the vacuum. Be careful not to allow the support to dry too much, as this will cause particle collapse and irreversible damage to the pore structure.

2. In the fume hood, prepare 100 ml of a solution consisting of 80 mg/ml DSC in dry acetone. Stir vigorously to dissolve. There may be some insoluble DSC left in the solution as a fine suspension, but this material will be driven into solution as the activation reaction proceeds.

3. Add the acetone wet gel cake to the DSC solution with stirring to uniformly resuspend the support. To this suspension, add 6.5 g of DMAP (alternatively, add of 7.5 ml of anhydrous TEA) to provide a base to catalyze the reaction. Mix the gel suspension using an overhead stirring paddle or by end-over-end rocking in the fume hood.


acetone to thoroughly remove excess reactant and reaction byproducts. Wash with 5 to 10 bed volumes of solvent to ensure removal of the last traces of unreacted DSC.

6. Store the NHS carbonate-activated support as a 50% slurry (the volume of the support is equal to half the total volume of the slurry) in dry acetone at 4°C until use. As an alternative storage solvent, the activated gel may be washed into dry isopropanol and stored in this solvent. Avoid the use of methanol, as some transesterification may occur with the NHS esters to form methyl esters upon storage.

The activation of an amine-containing support using DSC can be performed similar to methods described in previous protocols, except for substitution of a support modified to contain a terminal amino spacer for the hydroxylic or carboxylate supports described above.

LIGAND COUPLING TO NHS ESTER- OR NHS CARBONATE-ACTIVATED SUPPORTS

The following protocols describe general methods for coupling amine-containing ligands to NHS ester- or NHS carbonate-activated chromatography supports. This process can be used successfully to immobilize proteins or other amine-containing affinity ligands as well as amine-containing spacer molecules. During the coupling reaction, avoid introducing any other pri-mary or secondary amine-containing components into the coupling buffer—such as Tris, glycine, imidazole, ammonium ions, or other small molecules containing a reactive amine—as these will interfere with the desired immobilization of ligand. Amine reactions with reactive ester supports will occur efficiently in aqueous solution



at pH 7 to 9.0, so the coupling buffer pH may be modi-fied somewhat to promote ligand stability or solubility if necessary; however, be aware that the higher the pH of the reaction, the greater the rate of hydrolysis of the NHS esters, so use care when reacting at the high end of this pH range. The immobilization reaction may also be done entirely in nonaqueous solution or in a combi-nation of organic solvent/aqueous buffered solution to promote ligand solubility, when necessary. Nonaqueous conditions will eliminate the potential for hydrolysis and thus promote higher yields when immobilizing small amine-containing ligands. To perform immobili-zations in 100% organic solvent, an organic base should be added as a proton acceptor to catalyze the coupling reaction, such as triethylamine (TEA), diisopropylethyl-amine (DIEA), or dimethylaminopyridine (DMAP).

(A) Aqueous Reaction Protocol1. Prepare an amine-containing ligand solution

consisting of a protein or small molecule dissolved in 100 ml of 0.1-M sodium phosphate, 0.15-M NaCl, pH 7.2 (coupling buffer), at a concentration of 1 to 20 mg protein/ml or 1 to 5 mg/ml of a small amine-containing molecule. Alternative coupling buffers for immobilization onto NHS-activated supports include 0.1-M MOPS (pH 7.0), 0.1- to 0.2-M phosphate (pH 7.2–7.5), 0.1 to 0.2-M NaHCO3 (pH 8.0), or 0.1-M sodium borate (pH 8.5), which all may or may not contain NaCl. The pH of the ligand solution may be varied from approximately pH 7 to 9 to take into account differential reactivities of some ligands. The optimal concentration of ligand that is used for the immobilization reaction should be determined experimentally to obtain the best performance of the affinity matrix after coupling. This can be done using only 1 to 2 ml of activated support for each immobilization trial and proportionally reducing the quantities of reagents used for the reactions. Diamine spacer molecules should be coupled at much higher concentrations to avoid internal crosslinking while promoting coupling to only one end of the linker (i.e., 0.5–1.0 M). See the section on spacer arm coupling in this chapter for additional information.

2. In a fume hood, transfer 100 ml of an NHS ester- or NHS carbonate-activated support stored in isopropanol or acetone to a clean sintered glass filter funnel in the hood positioned over a suction filter flask. Drain the matrix of excess solvent by applying a vacuum and break up the resin with a plastic spatula into small pieces as the solvent is removed, then remove the vacuum to prevent gel drying. Wash the resin quickly with 2 to 3 bed volumes of deionized water to remove the majority of remaining solvent. When working with small quantities of

activated support, this operation can be performed using spin columns for solvent removal and washings. Finally, wash the activated support with 100 ml of coupling buffer and drain to a wet cake. Perform all aqueous washing operations quickly to avoid extensive hydrolysis of the reactive groups prior to mixing with ligand solution.

3. Immediately add the washed support to the ligand solution with stirring to uniformly mix and suspend the resin within the medium. Continue mixing using an overhead paddle stirrer or by rotation using a sealed container. React for at least 1 h at room temperature. For sensitive ligands, all solutions can be cooled to 4°C and the reaction also performed in the cold; however, the reaction time should be at least doubled to accommodate a slower reaction rate. The majority of coupling will occur in the first 30 min at room temperature.

4. Upon completion of the reaction, transfer the support slurry to a clean sintered glass filter funnel suspended in a suction filter flask, drain excess ligand solution, and wash with several bed volumes of coupling buffer. The initial washes may be saved to determine the amount of ligand that coupled to the support. This is done by measuring the remaining ligand in washes and comparing this amount to the quantity initially reacted. The difference is the amount of ligand immobilized onto the support.

5. Block unreacted NHS ester groups by adding to the support an equal volume of 1-M ethanolamine in coupling buffer and mixing for 30 min. The amino end of ethanolamine will couple to the remaining NHS ester groups and leave hydrophilic hydroxyl groups on the support.

6. Wash the support extensively with water, 1-M NaCl, and water, using at least 5 bed volumes of wash for each solution. Store the immobilized ligand support at 4°C as a 50% aqueous slurry containing a preservative.

(B) Nonaqueous Coupling Protocol1. In a fume hood, dissolve an amine-containing ligand

in 100 ml of a dry organic solvent at a concentration of 1 to 5 mg/ml. Suitable solvents include acetone, dioxane, DMF, DMSO, DMAC, ethanol, and isopropanol. The use of methanol may also be done, but it could more easily than other alcohols undergo transesterification with the NHS ester-reactive groups and slow down or reduce the coupling yield. Other water-miscible but dry solvents may be used as long as the solvent does not harm or unduly collapse the particle structure of the activated chromatography support. Add to this solution a 2-times mole excess of an organic base over the



amount of ligand present, such as TEA, DIEA, or DMAP to catalyze the coupling reaction.

2. Drain 100 ml of an NHS ester-activated support of excess solvent by using a sintered filter funnel suspended in a suction filter flask in a fume hood. If the coupling reaction is to be done in a solvent other than the one in which the activated support was stored, then wash the matrix into the coupling solvent by initially pulling a vacuum on the support in the filter funnel while breaking up the resin into small pieces with a spatula. Do not let the support dry in the filter funnel during the process. Remove the vacuum and resuspend the matrix in an equivalent volume of the desired solvent. Then wash with several volumes of this solvent to fully exchange the matrix into it. Drain to remove excess solvent.

3. Add the activated support to the ligand solution with stirring to uniformly resuspend the resin. Mix in a fume hood by using a paddle stirrer or by rotation in a sealed container. Continue the reaction for 1 h at room temperature with constant mixing.

4. Transfer the gel reaction slurry to a clean sintered glass filter funnel and drain the excess ligand solution. Wash with several bed volumes of solvent to remove the majority of not-coupled ligand. Retain the washes to determine the amount of ligand that coupled to the matrix, if desired.

5. Block unreacted NHS ester groups by adding to the support an equal volume of 1-M ethanolamine in solvent and mixing for 30 min. The amino end of ethanolamine will couple to the remaining NHS ester groups and leave hydrophilic hydroxyl groups on the support.

6. Wash the support extensively with solvent to remove all traces of ethanolamine and any remaining uncoupled ligand (at least 5–10 bed volumes). Next, wash the support into aqueous solution by sequentially washing with increasing amounts of water in solvent until 100% water is used and all traces of solvent have been eliminated from the resin. The final affinity support may be stored at 4°C as a 50% slurry in water containing a preservative.

Carbonyl Diimidazole (CDI) ActivationN,N’-Carbonyl diimidazole (CDI; also called 1,1′-car-

bonyl diimidazole) is a highly reactive carbonylating compound originally developed for use in peptide syn-thesis, but which is also capable of activating carboxyl-ates and hydroxyls on chromatography supports for the immobilization of amine-containing ligands (Paul and Anderson, 1960; Bartling et al., 1973; Bethell et al., 1979, 1987; Hearn et al., 1981, 1983; Batista-Viera et al., 2011). CDI is extremely sensitive to hydrolysis in water, where it decomposes to give two molecules of imidazole

with loss of CO2. Placing CDI in an aqueous solution will result in vigorous fizzing as the CO2 is released (Figure 15.48). All activation procedures, therefore, must be done in nonaqueous conditions using extremely dry solvents. Typically, water-miscible solvents are used so that the support can be washed in and out of aqueous solution by exchange with increasing or decreasing con-centrations of the solvent in water. Solvents that work well in this process include acetone, dioxane, DMSO, DMF, and DMAC. Avoid alcohols such as methanol, ethanol, or isopropanol, as the hydroxyls on these com-pounds will react with CDI.

N N

O

N N

CDIN,N '-Carbonyl diimidazole

The activation of a carboxylate group with CDI pro-ceeds through transamidination to give a secondary amide with imidazole. This intermediate acyl imidazolide is reactive towards amines in proteins and other mol-ecules, because the imidazole is a good leaving group. A coupling reaction involves attack of the nucleophilic amino group nitrogen on the carbonyl of the acyl imidaz-olide to displace the imidazole group and form an amide linkage with the ligand (Figure 15.49). Carboxylate-containing supports that can be activated with CDI include chromatography resins modified to contain a carboxylate-terminal spacer (see section on spacer arms, this chapter), membranes similarly modified with carbox-ylates, and small microparticles or nanoparticles that con-tain carboxylates (see Chapter 14, Section 4.2).

Hydroxyl-containing supports can also can be acti-vated using CDI, including those that contain primary or secondary hydroxyls (Hearn et al., 1983). In this case, CDI can be used to activate the hydroxyls to reactive imidazolyl carbamates, an intermediate that is similar in its reactions to NHS carbonates (described in the previ-ous section). Hydroxylic resins activated in this manner can then be used to immobilize amino ligands through displacement of the imidazole leaving group on the car-bonyl with subsequent formation of an alkyl carbamate linkage (Figure 15.50). The resulting urethane derivative

N N

O

N N

CDI

H2O

N

NH CO2+2

Imidazole

FIGURE 15.48 Hydrolysis of CDI in water to form CO2 and imidazole.



N N

O

N N

CDI


+

Acyl imidazole-activated support

Nonaqueous

H2N


Ligand immobilizedvia amide bond

Aque

ous

orno

naqu

eous

O

OH

O

N HN

N N+ + CO2

O

NH

FIGURE 15.49 Activation of carboxylates with CDI and reaction with amine-containing ligands.

N N

O

N N

CDI

OH


+

O

Imidazolyl carbamate-activated support

Nonaqueous

H2N


Ligand immobilizedvia carbamate bond

Aque

ous

orno

naqu

eous

O

N

N+

O

O

NH

N

NH

FIGURE 15.50 Activation of hydroxyls with CDI and reaction with amine-containing ligands.



creates an uncharged bond on the support that has excellent chemical stability toward ligand leakage. While both the activation of carboxylates and the acti-vation of hydroxyls with CDI produce an intermediate reactive carbonyl derivative, the respective bonds they form with amines are different, as are the hydrolysis products that are formed if a ligand does not couple in aqueous solution. If a CDI-activated carboxylate under-goes hydrolysis instead of ligand coupling, the result is a reformation of the original carboxylic acid group on the support. However, if a CDI-activated hydroxyl group is hydrolyzed, the result is loss of imidazole and CO2 along with reformation of the original hydroxyl. Thus, if a carboxylate support that has been activated with CDI should hydrolyze during a coupling reaction, it can create negatively charged groups on the matrix, which may cause subsequent nonspecific binding dur-ing affinity separations; whereas, if a hydroxylic support activated with CDI undergoes hydrolysis, it reverts back to a hydrophilic and uncharged hydroxyl, which helps to reduce nonspecific interactions (Figure 15.51). These reactions and their hydrolysis products are similar to the activation of hydroxylic- or carboxylic-containing sup-ports with DSC (described previously).

CDI-activated carboxylate supports or CDI-activated hydroxyl supports both will react nearly exclusively with primary amines on proteins and other ligands, even in the presence of secondary amines (Rannard and Davis, 2000). This surprising specificity can be used to advantage during the immobilization of spacer arms or small ligands, which may contain both primary and secondary amines within their structure. This may also be an important consideration in certain ligand immo-bilization needs in choosing CDI activation over the use of NHS ester- or NHS carbonate-activated supports, because they have the potential to react with secondary amines as well as primary amines.

Amine-containing supports may also be activated with CDI to give an intermediate imidazolyl isourea derivative. This reactive group also can be coupled with primary amines on ligands to result in an isourea (or urethane) bond (Figure 15.52). While the activa-tion of amine-containing supports is less frequently performed than the activation of carboxylate- or hydroxyl-containing supports, it does open up an additional route of activation and ligand coupling that avoids having to convert the amine to some other func-tionality prior to activation. The only potential defi-ciency of this method is the fact that amine-containing supports typically have greater nonspecific binding with biomolecules due to their net positive charge at physiological pH. Hydrolysis rather than ligand cou-pling of the CDI-activated intermediate would cause a reversion to a primary amine on the support, which would be the major cause of a positively charged affin-ity support.

Hydroxylic supports (such as agarose, Trisacryl, or Toyopearl matrices) that are activated with CDI and stored in nonaqueous solvent are stable for years if kept in a sealed container at 4°C. In addition, the sta-bility of the imidazolyl carbamate reactive groups is much greater toward hydrolysis in aqueous coupling buffer than NHS ester- or NHS carbonate-activated supports. At pH 8.5 to 9.0 in sodium borate buffer, a CDI-activated agarose support can take up to 30 h at room temperature to completely lose activity due to hydrolysis. By comparison NHS ester or NHS carbonate supports have half-lives due to hydrolysis measured in minutes in this pH range.

The immobilization of amine-containing ligands onto CDI-activated supports can be performed in non-aqueous or aqueous environments. It is particularly advantageous to couple small, organic soluble ligands in 100% organic solvent, as it eliminates the potential

O

Imidazolyl carbamate-activated support

O

N

N

O

N

N

H2O

N

NH+

H2O

Acyl imidazole-activated support

ImidazoleCarboxylate support

O

O–

N

NH CO2+

ImidazoleHydroxyl support

OH +

FIGURE 15.51 Hydrolysis of CDI-activated hydroxylic supports and CDI-activated carboxylate supports to give hydroxyls and carboxyls, respectively.



for hydrolysis during the reaction, thus maximizing the coupling yield. For aqueous-phase reactions, as in the coupling of proteins or other amine-containing bio-molecules, the optimal pH for immobilization often is dependent on the pI or pKa of the ligand. For maximal efficiency of coupling, the reaction conditions should be controlled at a pH representing at least 1 pH unit above the ligand’s pI or pKa point. This ensures that a major-ity of amines on the ligand will be unprotonated and thus at their maximal potential nucleophilicity for reac-tion with the active groups on the matrix.

CDI ACTIVATION PROTOCOL

The following protocols describe the activation of supports using CDI followed by a number of coupling methods that can be used to immobilize affinity ligands. Regardless of the support being used, the activation reaction should be done in anhydrous organic solvent to avoid hydrolytic breakdown of CDI or the active intermediate formed on the matrix. Make certain to use extensive solvent washing protocols to completely remove any traces of water from a hydrated support prior to the addition of CDI. Finally, when working with solvents all operations should be carried out in a fume hood using personal protective equipment and precautions to avoid static electricity. Dispose of all sol-vents according to hazardous organic waste guidelines.

1. Wash 100 ml of a hydroxyl-, carboxyl-, or amine-containing chromatography support material (such as crosslinked agarose) with water (at least several bed volumes) to remove preservatives and storage solution components. In a fume hood, sequentially wash the support into increasing concentrations of acetone-in-water (such as 25% (v/v), 50%, and 75% acetone/water) and finally with 100% dry acetone for at least 10 to 20 bed volumes to remove the last traces of water. Drain the support to a wet gel cake and then pull a gentle vacuum on the filter funnel while breaking up the gel cake with a spatula to obtain moist, evenly divided pieces, which still contain acetone within the internal pores of the beads. Be careful not to allow the support to dry, as this will cause particle collapse and irreversible damage to the pore structure of some matrices. Remove the vacuum as soon as the support is fully divided into small pieces (it will look like moist, fluffy snow). Note: Alternative solvents can be used for the activation process including dioxane, DMF, DMAC, or DMSO, which can be substituted for acetone depending on the compatibility of the matrix to a particular solvent. For example, some supports such as dextran will maintain their swollen particle nature best in solvents such as DMSO or DMF, whereas acetone will cause them to

N N

O

N N

CDI

NH2

Suppport containingamine groups

+

NH

Nonaqueous

H2N


Ligand immobilizedvia isourea bond

Aque

ous

orno

naqu

eous

O

N

N+

N

NH

NH

O

NH

Activated support Imidazole

FIGURE 15.52 CDI activation of amine-containing supports and coupling to amine ligands.



unacceptably shrink and restrict access to the inner pore structures.

2. In the fume hood, dissolve 10 g of CDI in 100 ml of dry acetone with stirring.

3. Add the acetone wet gel cake to the CDI solution with stirring to uniformly resuspend the support. Mix the gel suspension using an overhead paddle stirrer or by end-over-end rocking in a sealed container within the fume hood.


acetone to thoroughly remove excess reactant and reaction byproducts. Wash with at least 10 bed volumes of solvent to ensure removal of the last traces of unreacted CDI.

6. Store the CDI-activated support as a 50% slurry in dry acetone at 4°C until use.

Using the above protocol, the CDI-activated support may contain as much as 100 μmoles of reactive groups per milliliter (using crosslinked agarose), which is more than sufficient for coupling primary amine-containing affinity ligands of all types at very high density, if required.

LIGAND COUPLING TO CDI-ACTIVATED SUPPORTS

The following ligand coupling protocols represent generalized methods that should be optimized for a particular ligand to obtain the best possible affin-ity support performance in its intended application. CDI-activated supports typically have slower reac-tion kinetics when coupling proteins as compared to NHS ester-activated supports and therefore extended reaction times often are necessary to realize the best coupling yields (i.e., 16–24 h). However, for coupling small amine-containing ligands or spacer molecules, maintaining a high concentration should give maximal yields within 1 to 2 h at room temperature. Take care, however, in creating an immobilized ligand at very high density on the support, because this may cause nonspecific binding or result in too high an affinity toward the target molecule, making it difficult to elute it from the support.

For ligands that are more soluble or stable in organic solvent, the entire coupling reaction can be performed under nonaqueous conditions using the same sol-vent used in the activation process, if appropriate. If a nonaqueous reaction is performed, the addition of an organic base (e.g., TEA, DIEA, DMAP) at a concentra-tion that is 2 to 3 times that of the CDI activation level on the matrix will catalyze the process and increase the reaction rate. Reactions performed in organic sol-vent have the advantage of not undergoing hydrolysis, thus increasing potential yields of the ligand coupling

process. Alternatively, a mixture of aqueous buffer with solvent can be used to maintain the solubility of some ligands. In this situation, the ligand may be first dis-solved in organic solvent and then added to the reac-tion buffer just prior to adding the activated support. The final amount of solvent added to an aqueous/organic solvent reaction is often determined by the per-centage of solvent that can be tolerated by the buffered solution before buffer salts begin to precipitate. The maximal level of organic solvent addition can be dis-covered by making a series of solutions with increas-ing amounts of organic solvent to an aqueous coupling buffer and then identifying which concentrations do not cause buffer precipitation.

(A) Aqueous Coupling Protocol1. In a fume hood, drain 100 ml of CDI-activated

support of solvent using a sintered glass filter funnel suspended in a suction filter flask. Gently pull a vacuum to remove most of the excess solvent while breaking up the support into small, finely divided pieces, but be careful not to allow the matrix to dry out. Stop using suction as soon as the support is divided into small pieces. When coupling ligands that might precipitate or be damaged by the presence of some residual solvent, the support should be quickly washed using several bed volumes of cold deionized water to remove the organic phase. Drain the support to a wet gel cake.

2. Dissolve the ligand to be coupled in 100 ml of a buffered solution at pH 8.5 to 11, or in a buffer having a pH at least 1 pH unit above the pKa or pI of the ligand or protein. Proteins may be coupled at a concentration of 1 to 20 mg/ml or, for small molecules, at a concentration of 1 to 5 mg/ml of an amine-containing ligand. The optimal concentration of ligand that is used for the immobilization reaction should be determined experimentally to obtain the best performance of the affinity matrix after coupling. Suitable coupling buffers include 0.1-M sodium borate, pH 8.5 or 0.1- to 0.5-M sodium carbonate, pH 9 to 11. Avoid buffers or solution additives that contain amines, such as Tris, glycine, or imidazole, as these will compete with the ligand coupling reaction.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for at least 24 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. When using conditions at a pH of less than 9, the reaction time should be extended to 30 h to obtain maximal coupling yields. The reaction may be performed at 4°C or at room temperature, depending on the stability of the ligand or protein being immobilized. Excess CDI-reactive groups



on the support may be blocked by the addition of ethanolamine to the reaction slurry at a final concentration of at least 0.1-M. Note: Titrate the ethanolamine solution to the proper pH by addition of HCl before adding it to the reaction solution. Continue to mix for 1 h at room temperature.

4. Filter and wash the affinity support to remove uncoupled ligand and reaction byproducts using coupling buffer, water, 1-M NaCl, and again with water. Depending on the ligand being coupled, other wash solutions may be used to completely remove unreacted ligand, such as detergents, denaturants, and high or low pH conditions. Finally, store the affinity support in water containing a preservative at 4°C until used.

(B) Nonaqueous Coupling Protocol1. In a fume hood, drain 100 ml of CDI-activated

support of excess acetone using a sintered glass filter funnel suspended in a suction filter flask.

2. Dissolve the amine-containing ligand to be coupled in 100 ml of acetone (or another water-miscible solvent) at a concentration of 1 to 5 mg/ml, which is normally sufficient for the immobilization of a small organic compound. The optimal concentration of ligand to be used in the coupling reaction should be determined experimentally on small quantities of activated support to identify the best affinity support performance in its intended application. Add an organic base to the ligand solution, such as DMAP, DIEA, or TEA, to make a final concentration of 2-mM.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for 1 to 2 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. Longer reaction times may be used if appropriate. Excess CDI-reactive groups on the support may be blocked by the addition of ethanolamine to the reaction slurry at a final concentration of at least 0.1 M. Continue to mix for 1 h at room temperature.

4. Transfer the gel slurry to a sintered glass filter in the fume hood that is suspended in a suction filter flask and wash extensively (at least 10 bed volumes) with solvent to remove the remaining ligand and reaction byproducts. If the ligand is detectable, continue to wash the support until no further ligand is detected in the washings. Finally, drain the support of excess solvent by pulling a gentle vacuum on the filter flask while breaking up the support into small, finely divided pieces using a spatula, but be careful not to allow the matrix to dry out. Once the support is broken into small pieces, remove the vacuum and resuspend the gel in water with mixing. Continue

to wash the support with water until all traces of solvent have been removed. Additional washes with 1-M NaCl as well as low and high pH conditions may be carried out as appropriate. Finally, wash with water and store the affinity support as a 50% slurry in water containing a preservative at 4°C until used.

Activation Using FMP (2-Fluoro-1-Methylpyridinium)

Ngo in 1986 first described the use of 2-fluorometh-ylpyridinium (FMP), as the toluene-4-sulfonate salt, in a simple process to activate primary and secondary hydroxyls on supports and surfaces for the coupling of amine- or thiol-containing ligands. Under nonaqueous conditions, FMP reacts with hydroxyl groups on chro-matography supports to form an intermediate reactive methylpyridinium ether group, which can be coupled to ligands in aqueous or nonaqueous conditions to give stable secondary amine or thioether linkages. The immobilization reaction occurs in buffered solutions within the pH range of 5 to 10, but with greater reaction rates observed in the higher pH range. The coupling of ligands may also be performed in dry solvents with the addition of an organic base to catalyze the reaction (e.g., TEA, DIEA, DMAP) for ligands that are insoluble in aqueous solution.

The leaving group in the FMP coupling reaction (due to nucleophilic displacement with amino- or thiol-containing ligands) is 1-methyl-2-pyridone, which is a convenient chromophore that can be used to deter-mine the activation level of an FMP-activated support. The molar extinction coefficient of this leaving group is 5900 M−1 cm−1 at 297 nm, which also permits the cou-pling reaction to be followed provided the reaction is performed under nonaqueous conditions to eliminate simultaneous hydrolysis. Even in aqueous buffers, FMP-activated supports display relatively low hydrolysis rates. In acidic conditions, the activated support hydro-lyzes extremely slowly and even after months of stor-age retains a significant number of reactive groups. Ngo (1988) found that in 10-mM phosphoric acid, an FMP-activated support stored at 4°C for months was still able to couple 15 mg/ml of BSA. Even under alkaline condi-tions the activated support hydrolyzes slowly with a half-life of about 130 h at pH 7 and about 35 h at pH 9. Long-term storage in aqueous solution, however, still is not recommended, because some reactive groups will degrade over time and the reproducibility of coupling ligands may be negatively affected. One benefit of FMP-mediated immobilization is that if the activated support does hydrolyze during ligand coupling, the result is to reform the original hydroxyls on the support, thus leav-ing behind no groups that potentially can cause non-specific binding. In addition, successful ligand coupling



displaces the pyridone group, which bears the oxygen originating from the hydroxyl on the support material. Therefore, the net result of the immobilization reaction is to replace the hydroxyl group on the matrix with the nucleophile of the ligand, forming a new, chemically sta-ble bond (Figure 15.53). This process is similar to that of tosyl and tresyl activation, which also replaces the acti-vated hydroxyl with the amino group of the ligand (see tosyl and tresyl activation discussion, this chapter).

Amine- or thiol-containing ligands react with the FMP-activated support within the first several hours to result in at least 90% yields with regard to the amount of reactive groups that can react. Thiols appear to react at slightly greater rates due to their somewhat greater nucleophilicity. FMP-activated agarose or Toyopearl type supports should be able to immobilize proteins such as BSA at densities of 10 to 20 mg/ml gel.

After the coupling reaction is complete, the residual reactive groups should be blocked with a small molecule to ensure that no further reactivity remains that might couple molecules nonspecifically from subsequent sam-ples applied to the affinity gel. This blocking step is espe-cially important with FMP-activated matrices, because of the extended hydrolytic stability of the reactive groups. Blocking with ethanolamine, for example, will remove these active groups and result in numerous hydrophilic hydroxyls being created in their place.

FMP ACTIVATION PROTOCOL

The following protocols describe two methods of activating hydroxyl-containing supports with FMP in nonaqueous environments. The first one uses acetone and DMF as the solvents and the second method uses acetone and acetonitrile. Other solvents may be used for the activation step, but they should be water mis-cible so that water can be removed from the support by washing and anhydrous to prevent hydrolysis of the activation agent before the hydroxyls are activated.

(A) ACTIVATION OF TOYOPEARL SUPPORTS WITH FMP IN DMF

1. Wash 100 ml of a Toyopearl resin (such as HW-65 or HW-75) with water (at least several bed volumes) to remove preservatives and storage solution components. In a fume hood, sequentially wash the support into increasing concentrations of acetone-in-water (such as 25% (v/v), 50%, and 75% acetone/water) and finally wash with 100% anhydrous acetone for at least 10 to 20 bed volumes to remove the last traces of water. Drain the support to a wet gel cake and then pull a gentle vacuum on the filter funnel while quickly breaking up the gel cake with a spatula to obtain moist, evenly divided pieces, which still contains acetone within the internal pores of the beads. Be careful not to allow the support to dry. Remove the vacuum and add 200 ml of anhydrous DMF with stirring. Filter to remove excess DMF and then further wash the support with 500 ml of DMF to remove the acetone. Finally, filter off the excess solvent and keep as a moist gel cake.

2. In the fume hood, dissolve with stirring 7.1 g of FMP (5 mmoles; as the toluene-4-sulfonate salt) in 100 ml of dry DMF containing 6 mmoles TEA (3.03 g or 4.18 ml) as catalyst. Alternative organic base catalysts may be used, including DIEA or DMAP, which serve to accept protons during the course of the activation reaction.

3. Add the DMF wet gel cake to the FMP solution with stirring to uniformly resuspend the support. Mix the gel suspension using an overhead paddle stirrer or by end-over-end rocking in a sealed container within the fume hood.


500 ml of DMF to thoroughly remove excess reactant and reaction byproducts. Filter off excess DMF under a gentle vacuum and break up the gel cake into small pieces with a spatula. Resuspend the gel in acetone and then wash with at least 10 bed volumes of acetone.

6. Store the FMP-activated Toyopearl support as a 50% slurry in dry acetone at 4°C until use.

FMP

OH


+O

Activated support

Nonaqueous

H2N


Ligand immobilizedvia secondary amine bond

Aque

ous

orno

naqu

eous

NH

NF

NCH3

CH3

TEA

HON+

+

+

CH3

1-Methyl-2-pyridone

+

FIGURE 15.53 Activation of a hydroxylic support with FMP and coupling of amine- or thiol-containing ligands.



(B) ACTIVATION OF CROSSLINKED AGAROSE SUPPORTS WITH FMP IN ACETONITRILE

1. Wash 100 ml of a hydroxyl-containing, crosslinked agarose support (such as Sepharose CL-4B) with water (at least several bed volumes) to remove preservatives and storage solution components, then sequentially wash the gel in a fume hood with increasing concentrations of acetone-in-water (25% (v/v), 50%, and 75% acetone/water; 500 ml each). Finally, wash with 100% anhydrous acetone for at least 10 bed volumes to remove the last traces of water. Drain the support to a wet gel cake and then wash with at least 500 ml of dry acetonitrile. Finally, filter off the excess solvent and keep as a moist gel cake.

2. In a fume hood, dissolve 7.1 g of FMP (toluene- 4-sulfonate salt) in 300 ml of acetonitrile containing 4.18 ml of TEA.

3. Add the FMP solution to the moist gel cake and stir to uniformly mix the suspension.

4. Continue to react with mixing for 1 h at room temperature using an overhead paddle stirrer or end-over-end rocker.

5. In the fume hood, wash the activated support with 500 ml of acetonitrile to thoroughly remove excess reactant and reaction byproducts. Filter off excess acetonitrile under a gentle vacuum and break up the gel cake into small pieces with a spatula. Resuspend the gel in acetone and then wash with at least 10 bed volumes of acetone to remove the acetonitrile.

6. Store the FMP-activated agarose support as a 50% slurry in dry acetone at 4°C until use.

LIGAND COUPLING TO FMP-ACTIVATED SUPPORTS

The following immobilization protocols are general-ized methods for coupling amine- or thiol-containing affinity ligands to FMP-activated chromatography sup-ports. The optimal protocol for a given affinity ligand should be discovered through experimentation using different coupling conditions and concentrations of ligand to produce the best-performing affinity sup-port in the intended application. The first procedure is an aqueous coupling method that is appropriate for immobilizing proteins or small water soluble mole-cules. In the second procedure, a nonaqueous method is described that can be used to maximize the coupling yield (without competing hydrolysis) or for use with ligands that are not soluble in aqueous buffers.

(A) AQUEOUS COUPLING PROTOCOL

1. In a fume hood, drain 100 ml of the FMP-activated support of solvent using a sintered glass filter funnel suspended in a suction filter flask. Gently pull a vacuum to remove most of the excess solvent while breaking up the support into small, finely divided

pieces using a spatula, while also being careful not to allow the matrix to dry out. Stop using suction as soon as the support is divided into small pieces. When coupling ligands that might precipitate or be damaged by the presence of some residual solvent, the support should be quickly washed using several bed volumes of deionized water to remove the majority of acetone still present within the gel. Drain the support to a wet gel cake.

2. Dissolve the ligand to be coupled in 100 ml of a buffered solution at pH 7 to 9. As a guideline, proteins may be coupled at a concentration of 1 to 20 mg/ml or for small molecules, at a concentration of 1 to 5 mg/ml for an amine-containing ligand (or thiol-containing ligand). The optimal concentration of ligand that is used for the immobilization reaction should be determined experimentally to obtain the best performance of the affinity matrix after coupling. Suitable coupling buffers include 0.1 M sodium borate, pH 8.5, or 0.1- to 0.5-M sodium carbonate, pH 8.5 to 9. More physiological coupling conditions may also be used, such as 0.1-M sodium phosphate, pH 7.5, especially for molecules sensitive to higher pH conditions. Avoid buffers or solution additives that contain amines or thiols, such as DTT, 2-mercaptoethanol, glutathione, Tris, glycine, or imidazole, as these will compete with the ligand coupling reaction.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for at least 2 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. For ligands that have slower reaction kinetics with the FMP reactive group, the reaction time may be increased to overnight. In addition, when using conditions at a pH of less than 9, the reaction time can be extended to as much as 30 h to obtain maximal coupling yields. The reaction may be carried out at 4°C or at room temperature, depending on the stability of the ligand being immobilized.

4. Filter and wash the affinity support to remove uncoupled ligand and reaction byproducts using coupling buffer, water, 1-M NaCl, and again with water. Depending on the ligand being coupled, other wash solutions may be used to completely remove unreacted ligand, such as detergents, denaturants, and high or low pH conditions. Finally, store the affinity support in water containing a preservative at 4°C until used.

(B) NONAQUEOUS COUPLING PROTOCOL

1. In a fume hood, drain 100 ml of the FMP-activated support of excess acetone using a sintered glass filter funnel suspended in a suction filter flask.



2. Dissolve the amine-containing ligand to be coupled in 100 ml of acetone (or another water-miscible solvent) at a concentration of 1 to 5 mg/ml, which is normally sufficient for the immobilization of a small organic compound. If a bifunctional spacer molecule is to be coupled to the activated gel, such as a diamine compound, then much higher concentrations should be used (i.e., 0.5–1.5-M) to avoid internal crosslinking of the support during the reaction. The optimal concentration of ligand to be used in the coupling reaction should be determined experimentally on small quantities of activated support to identify the best affinity support performance in its intended application. Add an organic base to the ligand solution, such as DMAP, DIEA, or TEA, to make a final concentration of 2-mM.

3. In a suitable vessel, mix the activated wet gel cake with the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for 1 to 2 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. Longer reaction times may be used if needed to obtain maximal yields, if appropriate.

4. Transfer the gel slurry to a clean sintered glass filter in the fume hood that is suspended in a suction filter flask and wash it extensively (at least 10 bed volumes) with solvent to remove the remaining ligand and reaction byproducts. Wash with considerably more solvent if the initial concentration of ligand or spacer molecule in the reaction medium was particularly high. If the ligand is easily detectable in the filtrates, continue to wash the support until no further ligand is detected in the washings. Finally, drain the support of excess solvent by pulling a gentle vacuum on the filter flask while breaking up the support into small, finely divided pieces using a spatula, but be careful not to allow the matrix to dry out. Once the support is broken into small pieces, remove the vacuum and resuspend the gel in water with mixing. Continue to wash the support with water until all traces of solvent have been removed. Additional washes with 1-M NaCl as well as low and high pH conditions may be done as appropriate to remove any noncovalently bound ligand molecules. Finally, wash with water and store the affinity support as a 50% slurry in water containing a preservative at 4°C until used.

Activation Using Organic Sulfonyl ChloridesOrganic sulfonyl chlorides are activating agents that

can facilitate the conjugation of hydroxyl-containing compounds to other nucleophiles, particularly amine-containing ligands. They have also been used in organic synthesis for many years (Tipson, 1944; Brown et al., 1967; Beard et al., 1973; Kabalka et al., 1986; Whitaker

et al., 2006) and were first applied to the immobilization of ligands onto hydroxylic chromatography supports by Nilsson and Mosbach (1980) (also see Nilsson and Mosbach, 1981, 1984; Nilsson et al., 1981). Perhaps the most well-known organic sulfonyl chloride compounds used for synthesis are 4-toluenesulfonyl chloride, which also is referred to as tosyl chloride or TsCl (or TosCl), and methanesulfonyl chloride (also called mesyl chlo-ride or MsCl). The sulfonyl chloride group of TsCl or MsCl can react with a hydroxyl or an amine to form a sulfonyl ester or a sulfonamide group, respectively (Carey and Sundberg, 2007) (Figure 15.54). The sulfo-nyl ester formed with hydroxyls is further reactive with other nucleophiles, such as amine-containing affinity ligands, to facilitate their covalent linkage to activated chromatography supports.

SO

OCl

Tosyl chloride(p-toluenesulfonyl chloride)

H3CH3C SO

OCl

Mesyl chloride(methane sulfonyl chloride)

Sulfonate esters that are formed from the activation of hydroxyl groups on chromatography supports with TsCl create particularly good leaving groups, which get displaced as an amine-containing ligand couples to the matrix (Figure 15.55). In this case, the reaction essen-tially converts the hydroxyl on the support to a sec-ondary amine derivative that bonds the ligand to the matrix. Tosylates also can react with other nucleophiles, such as with hydroxyl groups under higher pH con-ditions (as the alkoxide, RO−) to form ether linkages, with thiols (as the thiolate anion RS−) to form thioether bonds, and also under alkaline conditions with OH−, which causes hydrolysis back to the hydroxyl. Reaction with these groups in nonaqueous conditions requires the presence of an organic base to act as a proton accep-tor to catalyze the coupling.

H3C SO

OCl

Tosyl chloride

OH


+O

Tosyl-activated support

NonaqueousSO

OCH3

FIGURE 15.54 Activation of a hydroxylic matrix with TsCl to form reactive sulfonate esters.



The attack of a nucleophile on a sulfonate ester can occur through one of two reaction mechanisms, SN1 or SN2, the path of which may be governed by the sol-vent environment and the chemical groups immedi-ately adjacent to the ester (Cremlyn, 1996; James and Cremlyn, 2002). The normal site of nucleophilic attack is on the carbon atom attached to the sulfonyl ester, which causes cleavage of the ester with concomitant bond for-mation of the nucleophile to the carbon atom. However, it has been demonstrated that nucleophilic attack can also occur onto the sulfur atom of the sulfonyl ester, which in this case may cause displacement of the con-stituent attached to the sulfonate opposite to the ester. The result of this reaction is the formation of a sulfon-amide linkage with amine nucleophiles and retention of the sulfonate onto the originally activated hydroxyl group on the matrix (Figure 15.56).

Using TsCl or MsCl, the alternative reaction prod-uct consisting of a sulfonamide is an extremely minor side reaction that may not occur to any appreciable extent when immobilizing amine-con-taining ligands. Nevertheless, the reaction mecha-nism for coupling ligands using tresyl chloride

(2,2,2-trifluoroethanesulfonyl chloride) has been found to be quite different. Tresyl chloride was first used for organic synthesis reactions by Crossland et al. (1971) and later employed for the activation of agarose chro-matography supports by Mosbach and Nilsson (1981). It was Crossland et al. (1971) that demonstrated that the reaction of a tresyl ester with an amine compound resulted in the loss of 2,2,2-trifluoroethane sulfonate as the leaving group. This reaction process was believed to be true for tresyl-activated hydroxylic chromatog-raphy supports, as well (Nilsson and Mosbach, 1981). However, it subsequently was discovered that the actual reaction mechanism leads to the formation of a sulfonamide linkage on the matrix with an amine-containing ligand, not a secondary amine linkage as in the use of tosyl chloride coupling (Demiroglou and Jennissen, 1990; Demiroglou et al., 1994). Careful ele-mental analysis indicated that the sulfur atom remains behind and the fluorine atoms are released in the immobilization process. In addition, the reaction of a thiol-containing ligand was found to yield a thiosulfate ester bond, which can have avid binding characteristics toward some proteins in affinity separations, particu-larly antibodies (Hutchens and Porath, 1986; Scoble and Scopes, 1997; Hansen et al., 1998) (Figure 15.57).

SO

O

Cl

Tresyl chloride

CF3

The unique reaction mechanism of tresyl ester-acti-vated supports may be due to the presence and strong electron-withdrawing properties of the trifluoro group. This may create a more powerful electrophilic center at the sulfur atom than that normally produced at the car-bon atom when using tosyl ester-mediated activation and coupling. Thus, an attack by a nucleophile such as an amine would occur at the sulfur atom center of the sulfonate group instead of at the carbon atom immedi-ately adjacent to the ester, which in turn would cause displacement of the trifluoroacetyl group. Under basic conditions with an abundance of OH− ions, this group is further transformed after displacement within the reaction medium into acetic acid and three fluorine ions (F−). In addition, the strong electron-withdrawing effects of the trifluoroacetyl group results in faster reac-tion kinetics for tresyl ester-activated supports than for tosyl ester-activated ones.

Sulfonyl chloride activation and coupling has been used for the immobilization of numerous affinity ligands onto hydroxylic supports of all types. In addi-tion to its use in the activation of beaded chromatogra-phy supports, tosyl chloride has been used to activate partially hydrolyzed rayon/polyester cloth for the

O


SO

OCH3

Hydroxyl-containingligand

O

Ether linkage

HO

HN


Secondary aminelinkage

H2N

HS

Thiol-containingligand

S

Thioetherlinkage

Incr

easi

ng p

H

FIGURE 15.55 Reactions of active tosylates with amines, thiols, and hydroxyls.



coupling of antibodies and other proteins for purifica-tion of target molecules (Boyd and Yamazaki, 1993) as well as for the immobilization of Aspergillus ory-zae β-galactosidase onto cotton cloth (Albayrak and Yang, 2002) and for the general coupling of amine nucleophiles onto cellulose (Heinze et al., 2001). Even

relatively inert membranes, such as polypropylene, have been modified using oxygen-plasma treatment to produce –OH functionalities and then activated with tosyl chloride for the coupling of peptides (Gérard et al., 2011). In addition, tresyl chloride activation has been used to immobilize fibronectin onto titanium solid phases (Hayakawa et al., 2003) and to couple antibodies onto PEG-modified particles for use in immunoassays (Chen et al., 2009).

Still another alternative sulfonyl chloride compound to that of TsCl or tresyl chloride has been described for activation of chromatography supports. The reagent 4-fluorobenzenesulfonyl chloride (called fosyl chloride or fosCl) has been used to activate hydroxylic groups on polymeric supports for the coupling of proteins and other biomolecules (Chang et al., 1992). It was found that the strong electron-withdrawing properties of the fluorobenzene group provides enhanced reaction rates for the activation and coupling steps over that of tosyl chloride.

SULFONYL CHLORIDE ACTIVATION PROTOCOL

The following activation protocol is a general-ized procedure for the use of tosyl chloride to activate hydroxyl groups on beaded chromatography supports. Other sulfonyl chloride compounds may be used under similar conditions, such as tresyl chloride, mesyl chlo-ride, or fosyl chloride. The activated support may be used to couple amine-containing affinity ligands and proteins as well as thiol-containing and hydroxyl-con-taining molecules. All procedures should be carried out in a well-ventilated fume hood to avoid exposure

O

Tosyl-activatedsupport

SO

OCH3



HN

Secondary aminelinkage

O SO

ONH

Sulfonamidelinkage

Minorproduct

Majorproduct

H2N

H2N

FIGURE 15.56 Alternative routes of nucleophilic attack on sulfonyl esters.

OH

Support containinghydroxyl groups

+O

Activated support

Nonaqueous


Ligand immobilizedvia sulfonamide bond

Aque

ous

orno

naqu

eous

TEA

SO

O

Cl

Tresyl chloride

CF3

SO

OCF3

OS

O

OHN

H2N

FIGURE 15.57 Tresyl chloride activation and coupling reactions.



to potentially toxic or reactive compounds and solvent fumes. The use of a sintered glass filter funnel sus-pended in a suction filter flask can facilitate the wash-ing steps.

1. Wash 100 ml of a hydroxyl-containing chromatography support (such as crosslinked agarose, Toyopearl, or Trisacryl resin) with water (at least several bed volumes) to remove preservatives and storage solution components. In a fume hood, sequentially wash the support into increasing concentrations of acetone-in-water (such as 25% (v/v), 50%, and 75% acetone/water). Finally, wash with 100% dry acetone for at least 10 to 20 bed volumes to remove the last traces of water. Drain the support to a wet gel cake and then pull a gentle vacuum on the filter funnel while breaking up the gel cake with a spatula to obtain moist, evenly divided pieces, which still contains acetone within the internal pores of the beads. Be careful not to allow the support to dry, as this will cause particle collapse and irreversible damage to the pore structure of some matrices. Remove the vacuum as soon as the support is fully divided into small pieces (it will look like moist, fluffy snow). Note: alternative solvents can be used for the activation process including dioxane, DMF, DMAC, or DMSO, which can be substituted for acetone depending on the compatibility of the matrix to a particular solvent. For example, some supports such as dextran will maintain their swollen particle nature best in solvents such as DMSO or DMF, whereas acetone will cause them to unacceptably shrink and restrict access to the inner pore structures for activation. Whichever solvent is used for the activation reaction, it should be anhydrous to prevent decomposition of the sulfonyl chloride by hydrolysis.

2. In the fume hood, dissolve 11.75 g of tosyl chloride (61.6 mmoles) in 100 ml of dry acetone with stirring. If an alternative sulfonyl chloride activating agent is to be used, add an equivalent mole quantity to the acetone solvent, and dissolve with mixing.

3. Add the acetone wet gel cake to the tosyl chloride solution with stirring to uniformly resuspend the support. Add 123 mmoles of an organic base as a proton acceptor to catalyze the activation by taking up generated HCl from the sulfonyl chloride as it reacts with the hydroxyls on the support. Suitable bases include pyridine, triethylamine (TEA), diisopropylethylamine (DIEA), or dimethylaminopyridine (DMAP). Mix the gel suspension using an overhead paddle stirrer or by end-over-end rocking in a sealed container within the fume hood.

4. React for 1 h at room temperature with mixing.

5. In the fume hood, wash the activated support with acetone to thoroughly remove excess reactant and reaction byproducts. Wash with at least 10 bed volumes of solvent to ensure removal of the last traces of unreacted tosyl chloride.

6. Store the tosyl ester-activated support as a 50% slurry in dry acetone at 4°C until use.

LIGAND COUPLING TO SULFONYL CHLORIDE-ACTIVATED SUPPORTS

The following methods are generalized procedures for the immobilization of amine-containing ligands onto sul-fonyl chloride-activated supports. Nonaqueous reactions may be carried out for small organic molecules or spacer arms that are soluble in organic solvent. Aqueous reac-tions are more appropriate for the coupling or proteins or other biological molecules that are soluble and stable in a buffered medium. Thiol-containing or hydroxyl-contain-ing ligands may also be coupled to sulfonyl chloride-acti-vated supports using similar protocols. Thiol-containing molecules will typically react faster than amines and will require buffers at the lower end of the pH spectrum described below. Hydroxyl-containing ligands, however, will need conditions at the high end of the pH range, as they are less nucleophilic at lower pH values.

(A) AQUEOUS COUPLING PROTOCOL

1. In a fume hood, drain 100 ml of the sulfonyl ester-activated support of solvent using a sintered glass filter funnel suspended in a suction filter flask. Gently pull a vacuum to remove most of the excess solvent while breaking up the support into small, finely divided pieces, but be careful not to allow the matrix to dry out. Stop using suction as soon as the support is divided into small pieces. When coupling ligands that might precipitate or be damaged by the presence of some residual solvent, the support should be washed quickly using several bed volumes of cold deionized water to remove the organic phase. Drain the support to a wet gel cake.

2. Dissolve the ligand to be coupled in 100 ml of a buffered solution at pH 8.5 to 10. Proteins may be coupled at a concentration of 1 to 20 mg/ml or for small molecules at a concentration of 1 to 5 mg/ml of an amine-containing ligand. The optimal concentration of ligand that is used for the immobilization reaction should be determined experimentally to obtain the best performance of the affinity matrix after coupling. Coupling buffers that may be used in this reaction include 0.1-M sodium borate, pH 8.5, or 0.2- to 0.5-M sodium carbonate, pH 9 to 10. If the ligand to be coupled is stable at the higher alkaline pH conditions, then the use of 0.25-M sodium carbonate, pH 9.5 will provide excellent coupling yields. Avoid buffers or solution



additives that contain amines, such as Tris, glycine, or imidazole, as these will compete with the ligand coupling reaction. If a tresyl-activated support is being used to immobilize ligands, then the reaction can be done in a sodium phosphate buffer at pH 7.5 to 8.0 (such as 0.2-M sodium phosphate, pH 7.5), because the reaction rate is much greater even at lower pH values.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for 24 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. The reaction may be performed at at 4°C or at room temperature, depending on the stability of the ligand being immobilized. Note that the reaction rate using tresyl-activated supports should be much faster than tosyl reactions, thus maximal coupling levels may be reached within 2 to 4 h. The exact time of the coupling reaction should be optimized for the best performance of the affinity support being produced.

4. Filter and wash the affinity support to remove uncoupled ligand and reaction byproducts using coupling buffer, water, 1-M NaCl, and again with water. Depending on the ligand being coupled, other wash solutions may be used to completely remove unreacted ligand, such as washing with detergent solutions, denaturants, and high or low pH conditions. Finally, store the affinity support in water containing a preservative at 4°C until used.

(B) Nonaqueous Coupling Protocol1. In a fume hood, drain 100 ml of a sulfonyl

ester-activated support of excess acetone (or other solvent) using a sintered glass filter funnel suspended in a suction filter flask. Do not allow the support to dry.

2. Dissolve the amine-containing ligand to be coupled in 100 ml of acetone (or another water-miscible solvent) at a concentration of 1 to 5 mg/ml, which is normally sufficient for the immobilization of a small organic compound. The optimal concentration of ligand to be used in the coupling reaction should be determined experimentally on small quantities of activated resin to identify the best affinity support performance in its intended application. For organic solvent-based reactions, add an organic base to the ligand solution, such as DMAP, DIEA, or TEA, to make a final concentration of 2-mM.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for 1 to 2 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. Longer reaction times may be used if appropriate.

4. Transfer the gel slurry to a sintered glass filter in the fume hood that is suspended in a suction filter flask and wash extensively with at least 10 bed volumes of solvent to remove the remaining ligand and reaction byproducts. If the ligand is detectable, continue to wash the support until no further ligand is detected in the washings. Finally, drain the support of excess solvent by pulling a gentle vacuum on the filter flask while breaking up the support into small, finely divided pieces using a spatula, but be careful not to allow the matrix to dry out. Once the support is broken into small pieces, remove the vacuum and resuspend the gel in water with mixing. Continue to wash the support with water until all traces of solvent have been removed. Additional washes with 1-M NaCl as well as low and high pH conditions may be done as appropriate to remove any noncovalently bound ligand. Finally, wash with water and store the affinity support as a 50% slurry in water containing a preservative at 4°C until used.

Azlactone-Activated SupportsAn oxazolone is a heterocyclic anhydride that can be

prepared from an N-acyl amino acid derivative through a dehydration and cyclization reaction. A particular type of oxazolone, an oxazol-5(4H)-one, is also known as an azlactone and consists of a five-membered ring that contains nitrogen and oxygen within its hetero-cyclic structure (see structure). Oxazolones have been used for many years in organic synthesis and have also become important constituents within organic com-pounds as potential drug candidates for a variety of applications in medicinal chemistry (Rao and Filler, 1986; Bala et al., 2011). The creation of azlactone groups can be accomplished from carboxylic acids through the reaction of α-methyl alanine in a two-step process using a condensing agent as the catalyst. This process is a variant of the classic Erlenmeyer–Plöchl azlactone synthesis, which was described in the late 1800s to prepare oxazolones (Plöchl, 1884; Erlenmeyer, 1893). Under anhydrous conditions, a carboxylate will react with a condensing agent to form an activated interme-diate, which then goes on to react with the amine of α-methyl alanine to form an amide linkage (an N-acyl amino acid derivative). This intermediate can undergo a subsequent condensation reaction through cycliza-tion of the amino acid carboxylate end via dehydra-tion, which results in the azlactone functionality being formed (Figure 15.58). Suitable condensing agents to drive this reaction include anhydrides, alkyl chloro-formates, and carbodiimides in a nonaqueous environ-ment. A preferred cyclization agent is acetic anhydride, which can be used as the solvent to form the azlactone functional groups on a dry carboxylate-containing



support material. The reaction proceeds at 80 to 100°C by mixing for 2 h.

ON O

General structureof an azlactone

(oxazol-5(4H)-one)

The formation of similar cyclic azlactones on car-boxylate-containing chromatography supports can be used to create electrophilic reactive groups that can be exploited for the immobilization of amine-containing (nucleophilic) ligands. The creation of solid support materials containing azlactone reactive groups was first described by 3M Corporation (U.S. patents 4,871,824 and 4,737,560; Coleman et al., 1990; Hermanson et al., 1995). To eliminate the potential for nucleophilic attack on the azlactone C4 site, the amino acid used to form the azlactone should contain substitutions on the C4 carbon. A particularly convenient choice of amino acid in this regard is α-methyl alanine (or 2-dimethyl glycine). The two methyl groups prevent substitution at the C4 car-bon after ring formation and therefore direct all nucleo-phile coupling reactions at the C5 position, which results in the desired ring opening process combined with amide bond formation with an amine-containing mol-ecule (Figure 15.59).

Polymeric chromatography supports contain-ing azlactone groups may be created more conve-niently at the time of resin production through the use of an azlactone-containing vinyl monomer in a copolymerization reaction with other monomers. The reactive monomer vinyldimethyl azlactone (or 2-vinyl-4,4′-dimethylazlactone) has been used along with N,N’-methylene bisacrylamide as a crosslink-ing monomer to create polymeric beaded particles for affinity chromatography (Coleman et al., 1990; Stanek et al., 2005) (Figure 15.60). This route of polymer for-mation yields a reactive support immediately upon

manufacture without the need to create a carboxylate support first and then activate the carboxylates to azlac-tones. The degree of reactivity in such polymers may be modulated by controlling the ratio of the azlactone-containing monomer to the other non-functionalized monomer(s) present in the final support. Using this

R OH

OH2N

OH

O

OH

OR N

H

O

ON O

R

OH

OR N

H

O

+

Carboxylate alpha-Methylalanine

Amide bond formation

Condensingagent

Ringformation

2-Dimethylazlactone

FIGURE 15.58 Formation of an azlactone from a carboxylate and α-methyl alanine in the presence of a cyclization catalyst.

Suppport containingazlactone groups

+

Ring opening andamide bond formation


HN

O

N

O

NH

O

O

H2N

FIGURE 15.59 Reaction of an α-methyl alanine-containing azlac-tone with an amine.

NH

O

NH

O

N,N '-Methylene bisacrylamide

N

O O

Vinyldimethyl azlactone

+

NO

O

NO

O

NO

O

NO

O

NO

O

NO

O

NO

ON

O

ON

O

ON

O

ON

O

ON

O

O

HN

O

HN

O

NO

ON

O

ON

O

ON

O

ON

O

ON

O

O

NO

O

NO

O

NO

O

NO

O

NO

O

Crosslinked polymercontaining azlactone groups

FIGURE 15.60 Synthesis of azlactone beads from vinyldimethyl azlactone and methylene bisacrylamide.



method, particles have been made that contain from approximately 100 μmoles/ml of azlactone groups up to a level of about 300 μmoles/ml, depending on the percentage of vinyldimethyl azlactone used in the copolymerization process. This type of support was previously commercialized under the Emphaze name through 3M and Pierce, and it now is available under the UltraLink name from Thermo Scientific (Pierce).

Supports prepared with azlactone groups should be stored in anhydrous organic solvent or kept dried as a stabilized particle powder to prevent hydrolysis of the reactive groups. Although the hydrolysis rate of the azlactone ring is slow during coupling reactions com-pared to the rate of ligand immobilization, the contin-uous exposure of the activated support to an aqueous environment will still degrade the reactive groups over time. Drying supports often requires the addition of excipients to stabilize the particles to prevent irrevers-ible collapse or pore size damage. The UltraLink sup-port is dried from a solution containing a small amount of detergent to decrease surface tension and allow rapid rehydration within the internal pore structures of the beads upon the addition of the dry support to a reac-tion medium. Therefore, ligand coupling reactions can be performed simply by means of the measured addi-tion of the activated support to the appropriately buff-ered ligand solution and mixed to uniformly distribute the particles throughout the solution. Amine-containing ligands such as proteins can react with azlactone groups in a buffered environment over a pH range of 4 to 9, and even more extreme pH conditions may be used for coupling spacer arms or small organic mol-ecules that are stable to highly alkaline conditions. For instance, coupling a diamine spacer to an azlactone-activated support can be done at a concentration of 0.5 to 1.5-M without pH adjustment (pH >12) to accel-erate the reaction and increase immobilization densi-ties. Ligand coupling to the azlactone groups results in the formation of amide bonds with formation of a short spacer arm created from the ring opening pro-cess (Figure 15.59). A comparison of the coupling of protein A, protein G, protein A/G, avidin, and strepta-vidin using two different buffers at pH 7.5 and pH 9.0 was made by Hermanson et al. (1995). In general, higher coupling yields were observed at pH 9.0 for three dif-ferent ligand loading levels in each immobilization reaction.

An azlactone-activated support can have an initial degree of hydrophobic character due to the presence of the heterocyclic reactive groups. Many hydrophilic charged molecules such as proteins have difficulty approaching close enough to the activated support sur-faces to facilitate efficient coupling using reasonable reaction times. This problem can be overcome through the use of buffer additives that are lyotropic in nature

and which can have the effect of salting out and push-ing the proteins closer to the surfaces for coupling. Suitable lyotropic salts for this purpose can be cho-sen from the Hofmeister series, such as the addition of sodium sulfate to the coupling buffer at a concentra-tion of 0.8 to 1.5-M. The salt concentration should be adjusted to maintain protein solubility while maximiz-ing the rate of immobilization to the support. Using these conditions, the immobilization reaction can be complete within 1 h and achieve maximal yields. Protein A has been coupled to azlactone-activated sup-ports at densities of over 30 mg/ml and immunoglobu-lins have been immobilized to levels of over 20 mg/ml. After the coupling reaction is carried out, any excess reactive groups should be blocked using a small mol-ecule such as ethanolamine. Once an affinity support is prepared using these methods, the support material reverts to a hydrophilic surface with relatively low non-specific binding potential to the matrix itself.

Azlactone activation has been used for the cou-pling of affinity ligands to beaded chromatography supports (Coleman et al., 1990; Hermanson et al., 1995; Stanek et al., 2005), for the immobilization of ligands onto various polymeric constructs (Laquièvre et al., 2012), for modification and subsequent immobiliza-tion onto surfaces (Lokitz et al., 2009), and to couple enzymes to monolithic supports for use in microfluidic devices (Logan, 2007). Surface polymer scaffolds have also been built to immobilize affinity ligands using multilayer copolymers incorporating vinyldimethyl azlactone groups (Barringer et al., 2009). The use of the vinyl monomer is advantageous in creating copoly-mer constructs of many different structures as well as for photografting azlactone-reactive groups onto exist-ing materials for immobilization of amine-containing biomolecules.

LIGAND COUPLING TO AZLACTONE-ACTIVATED SUPPORTS

The following procedure describes a generalized method for the coupling of protein affinity ligands to 10 ml of an azlactone-activated beaded chromatography support. Similar methods may be used to immobilize biomolecules onto activated membranes, surfaces, or monolithic supports containing azlactone groups. Two additional procedures are subsequently presented that describe methods for coupling small amine-containing molecules, such as organic ligands and spacer arms, using aqueous or nonaqueous conditions.

(B) Protein Immobilization onto Azlactone Supports1. Dissolve the protein to be immobilized at a

concentration of 1 to 20 mg/ml in 20 ml of coupling buffer (sufficient for coupling to 10 ml



of swollen azlactone beads) consisting of 0.1-M buffer concentration and at a pH range of 4 to 9. Most proteins react best at the high end of the recommended pH range, but some optimization might have to be carried out to obtain the best coupling yield and affinity ligand performance for a particular protein to be coupled. Also add to the coupling buffer a lyotropic salt to enhance the rate of protein coupling to the azlactone-activated support. A suggested coupling buffer is 0.1-M sodium carbonate, pH 9.0, with the addition of 0.6-M sodium citrate as the lyotropic agent. An alternative lyotrope that may be used is sodium sulfate at a concentration of 0.8-M. A more neutral pH coupling buffer that has been used successfully is 0.1-M sodium phosphate at pH 7.5, containing 0.6-M sodium citrate or 0.8-M sodium sulfate. The concentration of the lyotropic salt may be lowered for proteins that have solubility issues at the recommended concentrations; however, the recommended levels for sodium citrate or sodium sulfate have been found to work well for antibody (IgG) immobilizations, for avidin or streptavidin, as well as for the coupling of immunoglobulin binding proteins such as protein A or protein G to an azlactone support. In addition, other buffer components may be used to stabilize the protein in the reaction medium at different pH levels, but avoid additives that contain amines or other nucleophiles that would compete with the reaction—for example, Tris, imidazole, thiol reducing agents, glycine, and ammonium ions. If the activated particles are added as a dry powder, prepare enough ligand solution to equal at least twice the volume of the hydrated beads. This volume is required because half of the ligand solution volume will be used to hydrate and fill the pore structure of the particles, while the other half will create a 50% slurry for efficient mixing. If the support is in a hydrated state prior to addition of the ligand solution, then prepare an equal volume to the amount of swollen support used.

2. Measure out a quantity of dry azlactone-activated particles corresponding to exactly half the volume of the ligand solution prepared in step 1—in this case, weigh out the equivalent of 10 ml of beads. For instance, a typical batch of UltraLink azlactone-activated beads has a swelling ratio of approximately 120 mg/ml hydrated gel (it varies slightly for each individual lot of manufactured particles, so refer to the lot-specific information to be precise). To yield 10 ml of hydrated beads, therefore, add 1.2 g of azlactone beads to the ligand solution with stirring to fully hydrate the particles and initiate the coupling reaction.

3. React with mixing for 1 to 2 h at room temperature using an overhead paddle stirrer or end-over-end

rotation in a sealed container. The majority of protein coupling reactions will reach maximal yield in 1 h.

4. Wash the support with several bed volumes of water to remove excess reactants by using a sintered glass filter funnel suspended in a suction filter flask. Note that if the UltraLink support is used, it is supplied dry from the manufacturer with a small amount of Triton X-100 added to promote rapid rehydration within the pores. The presence of the detergent in the initial washings after coupling will prevent the accurate measurement of protein concentration by absorbance at 280 nm. However, the protein concentration that did not couple to the support may be determined by use of the BCA assay, which is detergent tolerant (Smith et al., 1985). The measurement of the non-coupled protein in the washings can facilitate the determination of how much protein has coupled to the support by the difference between the amount of protein not coupled and the initial amount of protein used in the coupling reaction. Drain the support to a wet cake.

5. To block the remaining azlactone reactive groups, add 1 bed volume (10 ml) of 1-M ethanolamine, pH 9.0. Mix for 30 min. Note: Adjust the pH of the ethanolamine solution in a fume hood using the slow addition of 50% HCl while maintaining the solution on ice. Adjust to room temperature before using.

6. Wash the support with coupling buffer without containing the lyotropic agent, then wash extensively with 1-M NaCl and finally with water to completely remove unreacted molecules. Additional washes may be carried out using acid or basic pH conditions as well as employing the use of detergents or denaturants to ensure complete removal of any proteins still bound by noncovalent interactions. These additional wash conditions should be done only if the protein immobilized is stable to such environments and only if necessary to minimize the leaching of ligand molecules in subsequent chromatographic operations. Finally, store the affinity support in water or buffer containing a preservative at 4°C.

(B) Aqueous Coupling of Small Ligands or Spacers to Azlactone Supports

Affinity molecules may also be small organic com-pounds that can be immobilized onto an azlactone sup-port through an available amine group. Additionally, amine-containing spacer arms may be coupled to the support to facilitate further derivatization or to create a different chemical functionality at its terminal end for subsequent coupling to a ligand through some-thing other than an amine (e.g., a thiol). The aqueous coupling of small molecules to an azlactone support is performed somewhat differently from the coupling



methods typically used with proteins, because a lyo-tropic agent usually is not required to enhance the reac-tion rate and yield. Instead of using the salting out effect of a lyotrope to drive proteins toward the surface azlac-tone groups to enhance reaction rates, small-molecule coupling is improved through increased concentrations and higher pH conditions. Another caveat to consider is that if a molecule has more than one amine—as in diamine spacer arms, which makes multipoint attach-ment to the support more likely—then the concentra-tion of ligand used during the reaction should be very high (i.e., 1.0–2.0 M) to prevent internal crosslinking or even bead-to-bead linking caused by both ends of the compound reacting with the support. In this case, the high concentration range encourages single point attachments by making it more likely that another mol-ecule will react rather than a second site within the same molecule getting coupled. This is an especially important factor when coupling diamine spacers to any amine-reactive support, since the desired end product involves one amine getting immobilized and the other amine of the spacer remaining free for subsequent reac-tions. On the other hand, for spacer arms containing an amine on one end and another functional group, such as a carboxylate on the other end, the need for very high initial concentrations is not as important, since only one end of the molecule will be capable of coupling.

Coupling of Small Diamine Spacers1. In a fume hood, dissolve a diamine spacer molecule,

such as 3,3′-diaminodipropylamine (DADPA; also called bis(3-aminopropyl)amine), in water at a concentration of at least 1.5 M. When using diamines as the free base compounds (not as the hydrochloride salts), there is no need to adjust the pH once the solution is made. The pH will be extremely basic, but this will drive the coupling reaction to the azlactone groups on the support with higher efficiency (see additional information in the section on coupling spacer arms, this chapter). Prepare 20 ml of the diamine solution to react with 10 ml of hydrated azlactone beads.

2. Add 1.2 g of azlactone beads to the diamine solution with stirring to fully hydrate the particles and initiate the coupling reaction (this quantity is equivalent to 10 ml of particles once they are fully hydrated and swollen).

3. React for 1–2 h at room temperature with constant mixing by use of an overhead paddle stirrer or end-over-end mixing on a rotator (do not use a magnetic stirring bar).

4. Wash the amine-modified support using a sintered glass filter funnel in a fume hood. Wash extensively with water, 1-M NaCl, and again with water to remove excess diamine. Use 10 to 20 bed volumes of

washes for each solution. The presence of amines on the support, which indicates successful coupling, can be monitored by reacting a small quantity of support with TNBS solution, which will give a bright orange color with primary aliphatic amines (see Chapter 2, Section 4.3). Store the amine support in water or buffer containing a preservative at 4°C.

Coupling Small, Water Soluble Amino Ligands1. Dissolve an amine-containing ligand in 20 ml 0.1-M

sodium carbonate, pH 9.0, at a concentration that will result in the desired density of affinity groups on the surface, which ultimately will be optimal for the chromatography application. For some ligands, such as peptides, the concentration may be in the range of 3 to 5 mg/ml of support. If a high density of coupling is desired, the concentration of ligand may be increased to at least double the amount of azlactone groups on the surface (i.e., 2 × 100 μmoles/ml).

2. Weigh out 1.2 g of azlactone beads (10 ml gel after hydration) and add them to the ligand solution with stirring.

3. React for 2 h to overnight at room temperature with constant stirring.

4. If the ligand was reacted at a level that would not couple to every azlactone group present on the support surface, then wash the support with several column volumes of water to remove the majority of remaining uncoupled ligand. Next, add to the support 10 ml of 1-M ethanolamine, pH 9.0, to block the remaining reactive groups. React for 1 h at room temperature with mixing.

5. Wash the support with coupling buffer and then with water, 1-M NaCl, and again with water to remove uncoupled ethanolamine and ligand. Wash with at least 10 to 20 bed volumes of each solution. Store the amine support in water or buffer containing a preservative at 4°C.

(C) Coupling Amine Containing Ligands in Organic Solvent

Some amine-containing ligands may be only spar-ingly soluble in aqueous buffer conditions. For this rea-son, it may be necessary to perform the immobilization reaction in organic solution to maximize the coupling yields and obtain an optimal affinity support. The fol-lowing protocol describes an organic solvent reaction using azlactone particles (UltraLink). An important caveat is that the solvent chosen should maintain a swollen bead state so as not to restrict ligand access to the internal pore structures of the particles.

1. In a fume hood, dissolve an amine-containing ligand to be coupled in 20 ml of dry DMSO or DMF at a concentration of at least 2 to 5 mg/ml. The optimal



concentration of ligand in the immobilization reaction may have to be determined experimentally using small quantities of activated support and by varying the concentration of ligand to observe its affect on affinity chromatography performance. Other solvents may be used; however they should be water miscible and able to maintain a swollen bead state for the activated support.

2. Add 1.2 g of azlactone support (10 ml hydrated volume) to the ligand solution with stirring.

3. React for 1 to 2 h at room temperature with constant stirring.

4. Wash the support in a fume hood using a sintered glass filter suspended in a suction filter flask. Wash with at least 20 bed volumes of the organic solvent used for the coupling reaction. Continue to wash until no ligand is detected in the filtrate. Wash the support into water using progressively increasing concentrations of water in solvent (e.g., 30%, 70%) until 100% water is used. Wash with 100% water for at least 10 bed volumes to completely remove the last traces of solvent. Finally, store the affinity support as a 50% slurry in water or buffer at 4°C containing a preservative.

Cyanogen Bromide ActivationOne of the first activation methods introduced

for the coupling of affinity ligands to solid supports involves the use of cyanogen bromide (CNBr) to acti-vate hydroxyl groups (Axen et al., 1967). For many years, CNBr activation was the method of choice for coupling amine-containing ligands to agarose supports, espe-cially for the immobilization of proteins. Pharmacia first commercialized CNBr-activated agarose in a dry form under the name CNBr Sepharose and the product is still sold by GE for affinity ligand immobilization. Meng et al. (2009) recently reported on methods for stabilizing CNBr-activated agarose in dry form, which preserves

both the reactive group and the structure of the beaded support. Under alkaline conditions, CNBr reacts with the hydroxyl groups on a matrix to produce reactive cyanate esters and imidocarbonates (Figure 15.61). The relative yield of cyanate esters versus imidocarbonates may vary depending on the hydroxylic structure of the chroma-tography matrix. If closely spaced or adjacent hydrox-yls mainly are present in the support, such as within dextran- or cellulose-containing matrices that contain many diols, then a cyclic imidocarbonate may become the predominate product. However, if primary hydrox-yls or hydroxyls that are not immediately adjacent to one another are present within the support, such as in aga-rose matrices, then cyanate esters are the major product formed from the reaction (Kohn and Wilchek, 1982).

CNBr activation is highly versatile and has been used to activate hydroxyl groups on many different chromatography supports as well as on microparticles, nanoparticles, membranes, and even surfaces (Jurado et al., 2002; Yavuz et al., 2008; Arazawa et al., 2012). The major disadvantages of using this coupling method are the high toxicity of the reagent (it emits hydrogen cya-nide gas, which is deadly if inhaled), the potential for creating a positive charge on the support after ligand coupling due to the bond type that is formed (primar-ily an isourea linkage, which is positively charged at neutral pH), and the labile nature of the ligand linkage to the support (having a tendency to constantly leach immobilized ligands at levels higher than that observed using other coupling methods). The unfortunate disad-vantages of using CNBr activation have led to a signifi-cant decrease in its use over the years and an increase in the use of alternative amine coupling chemistries that do not have these shortcomings.

CNBr-activated supports react with amine-con-taining ligands to potentially produce two linkages, depending on whether the starting activation form was

OH


OH OH-

OH

O CNBr

OH

O C N

Cyanate ester(highly reactive)

O

O C NH

Cycliz

atio

n

Imidocarbonate(lower reactivity)

FIGURE 15.61 CNBr activation forming isocyanate and cyclic imidocarbonates.



a cyanate ester or a cyclic imidocarbonate. Cyanate esters rapidly react with amines under mildly alkaline pH conditions to form an isourea bond, whereas the cyclic imidocarbonate reactive group is far less reactive and couples with amines to create a substituted imido-carbonate linkage (Figure 15.62).

CNBr ACTIVATION PROTOCOL

The first procedure describes the original method reported by Cautrecasas (1970) of using titration with NaOH during the activation process to maintain the pH of the reaction. The second method describes the revised protocol of March et al. (1974), which involves reagent modifications that make the pro-cess easier to control. The reagent quantities may be proportionally scaled to activate different amounts of chromatography support materials. Read through the protocols thoroughly before setting up the activation reaction, because many of the steps are dependent on time, temperature, and pH, which all need to be controlled to result in successful and reproducible activation and coupling of an affinity ligand.

Caution: CNBr is highly toxic and may cause death if inhaled or ingested. Avoid contact with the solid com-pound and any solutions containing it. All operations should be carried out in a well-ventilated fume hood and using appropriate personal protective equipment. Dispose of all waste according to recommended safety protocols (see the product’s MSDS data sheet for further details).

(A) Traditional Method: CNBr Activation using NaOH Titration (Cuatrecasas, 1970)1. Wash the equivalent of 100 ml of settled

chromatography support containing hydroxyl groups with 1 l of deionized water using a sintered

glass filter funnel and a vacuum filter flask. Pull a gentle vacuum to facilitate the filtration process and collect all washes for proper disposal. After the wash, suction the support to a wet cake, stopping the filtration process by eliminating the vacuum at the point that the excess wash solution just enters the top of the gel. Do not allow the gel bed to get air within it or dry out at the top during the washing and filtering process. Note that if a monolithic support or a membrane is used in this procedure, then refer to the recommendations on handling these materials in Section 1 of this chapter.

2. Remove the washed and moist support from the filter funnel, transfer it to a beaker, and suspend it in 100 ml water by stirring. In a fume hood, set up an overhead paddle stirrer that will be used to mix the support during the activation procedure. Do not use a magnetic stir bar, as it will grind the support material and damage it. Insert a pH probe and a thermometer into the gel suspension to continuously monitor the pH and temperature during the activation reaction.

3. Prepare a 20% (w/w) solution of NaOH (i.e., 20 g/100 ml) by dissolving NaOH pellets in water or by dilution of a commercially available 50% solution. In addition, have available an ice bucket full of small, deionized ice chips to cool the reaction as it proceeds.

4. In the fume hood, weigh out 20 g of CNBr (caution: extremely toxic compound!) and add it to the stirring gel suspension. Maintain the pH of the reaction at pH 11 by dropwise addition of 20% NaOH solution. Also maintain the temperature of the reaction at about 25°C by periodic addition of crushed ice chips to the slurry.

5. Continue the activation reaction for 10 to 15 min at which point the CNBr should be completely

O C N

Cyanate ester(highly reactive)

O

OC NH

Imidocarbonate(lower reactivity)



Isourea linkage

Cyclic imidocarbonatederivative

OC

NH2

O

OC

NH

H2N

H2N

N

FIGURE 15.62 Ligand coupling to CNBr-activated supports.



dissolved and the rate of base consumption should be reduced. Transfer the activated support to a sintered glass filter funnel set up in the fume hood containing deionized ice chips to cool the reaction. Wash the support with 1 l of ice-cold deionized water followed by 500 ml of ice-cold 0.1-M sodium bicarbonate, pH 8.5 (coupling buffer). Drain the activated gel to a moist cake on the sintered glass filter pad.

6. Use the activated support to immediately couple an amine-containing ligand according to the immobilization protocol described below.

(B) Alternative Protocol: 2-min CNBr Activation using Carbonate Buffer at Room Temperature (March et al., 1974)1. Wash the equivalent of 100 ml of settled

chromatography support containing hydroxyl groups with 1 l of deionized water using a sintered glass filter funnel and a vacuum filter flask. Pull a gentle vacuum to facilitate the filtration process and collect all washes for proper disposal. Next wash the support with several bed volumes of 2-M sodium carbonate (no pH adjustment required). After the carbonate wash, suction the support to a wet cake, stopping the filtration process by eliminating the vacuum at the point that the excess wash solution just enters the top of the gel. Do not allow the gel bed to get air within it or dry out at the top during the washing and filtering process. Note that if a monolithic support or a membrane is used in this procedure, refer to the recommendations on handling these materials in Section 1 of this chapter.

2. Remove the washed and moist support from the filter funnel, transfer it to a beaker, and suspend it in 100 ml 2-M sodium carbonate buffer by stirring. In a fume hood, set up an overhead paddle stirrer that will be used to mix the support during the activation procedure. Do not use a magnetic stir bar, as it will grind the support material and damage it. Stir the gel slurry at a rate that keeps the particles well mixed and suspended in the activation buffer.

3. In a well-ventilated fume hood, weigh out 10 g of CNBr (caution: extremely toxic compound!) and dissolve it in 5 ml of acetonitrile.

4. Add the CNBr solution to the stirring gel slurry and react for exactly 2 min at room temperature.

5. Immediately transfer the gel to a sintered glass filter in the fume hood and wash with 1 l of ice-cold water followed by 500 ml of ice-cold coupling buffer (0.1-M sodium bicarbonate, pH 8.5). Drain the activated gel to a moist cake on the sintered glass filter pad.

6. Immediately use the activated support to couple an amine-containing ligand according to the following protocol.

LIGAND COUPLING TO CNBr-ACTIVATED SUPPORTS

The following protocol describes the general method for coupling amine-containing ligands to a CNBr-activated chromatography support material. This pro-cess can be used successfully to immobilize proteins or other amine-containing affinity ligands as well as amine-containing spacer molecules. Avoid introducing any other amine-containing components into the coupling buffer during the reaction, such as Tris, glycine, ammo-nium ions, or other small molecules containing a reactive amine, as these will interfere with the desired immobi-lization of ligand. Amine reactions with CNBr-activated supports will occur efficiently between pH 8 and pH 9.5, so the coupling buffer pH may be modified somewhat to promote ligand stability or solubility if necessary.

Protocol1. Suspend the CNBr-activated support in an equal

volume of 0.1-M sodium carbonate buffer, pH 8.5 (coupling buffer), into which an amine-containing affinity ligand has been dissolved. For many protein ligands, a suggested starting concentration is 3 to 6 mg/ml. For low-molecular-weight ligands use a concentration at least 3 times greater than the level of CNBr activation. For the activation of agarose with CNBr, the typical activation level is about 20 to 40 μmoles/ml gel.

2. Mix the reaction slurry using a paddle stirrer for 24 h at 4°C.

3. Using a sintered glass filter funnel, wash the gel extensively with coupling buffer, then with 1-M NaCl, and water to remove excess unreacted ligand, which did not get immobilized.

4. Excess reactive groups on the matrix can be blocked by the addition of 1-M ethanolamine, pH 9, or 1-M Tris, pH 9. Suspend the washed gel in an equal volume of the blocking solution and stir for 1 h at room temperature.

5. Remove excess blocking agent by washing the affinity support extensively with 1-M NaCl and water. The final washes can be tested for the presence of amines through reaction with TNBSA, which will turn orange in solution upon coupling to aliphatic amines (see Chapter 2, Section 4.3). Finally, store the affinity support as a 50% slurry in water or buffer at 4°C containing a preservative.

Trichloro-s-Triazine (TsT) ActivationCyanuric chloride or trichloro-s-triazine (TsT) is a

trifunctional, symmetrical, heterocyclic aromatic com-pound that has been used for many decades as a reac-tive group in the construction of dyes for staining fabrics and articles of clothing. Dyes such as the Procion series react through covalent bonding of the dye to the



nucleophilic groups on the fabrics to create stable, non-fading, colored clothes (see structure of Procion Brilliant Blue). TsT can also be used to activate and form reactive groups on the surfaces of hydroxylic chromatogra-phy resins for the immobilization of affinity ligands. The three reactive chlorines of TsT potentially can covalently link to three separate nucleophilic groups on different molecules. The reaction of TsT in excess with the hydroxyl groups on chromatography support materials can result in two reactive sites remaining for coupling of amine-containing ligands. However, the reactivity of the triazine chlorine groups on TsT varies depending upon how many have already reacted. The first chlorine is the most highly reactive, and it can cou-ple to the hydroxyls on supports under relatively mild conditions. The second reactive chlorine is able to cou-ple with a nucleophile such as an amine, thiol, or even a hydroxyl group under slightly basic pH conditions with relatively rapid reaction kinetics. Finally, the last chlo-rine of TsT has the lowest reactivity, but it still can be used to couple with thiols or amines under mildly alka-line pH conditions and with hydroxyls under highly alkaline conditions. TsT activation has also been used to conjugate PEG molecules to proteins for the modulation of immunological properties (Abuchowski et al., 1977) (Chapter 18, Section 2).

O O

H2N NH

S–O

O

O

SO–

O

ONH

N

NN

Cl

Cl

Procion Brilliant Blue

TsT activation of a hydroxyl-containing support such as agarose proceeds through loss of one chlo-rine (as HCl) with modification to the matrix through a hydroxyl by means of an ether linkage to the ring (Figure 15.63). If TsT is used in large excess, most of the active groups on the support will contain two remain-ing chlorines for coupling to affinity ligands. The activation process is best done under nonaqueous con-ditions to prevent hydrolysis of the TsT chlorines, and also while in the presence of two equivalents of organic base (e.g., DIEA) to accept the released protons pro-duced during the process (Finlay et al., 1978; Hodgins et al., 1980). TsT-activated supports represent one of the few truly multi-purpose reactive groups—others being epoxides, iodoacetyl, and vinyl sulfones—that are able to couple with amine-, thiol-, and hydroxyl-containing

ligands with success just by modulating the pH. In addition to the trichloro derivatives of triazine rings, the trifluoro derivatives have also been used with suc-cess for activation of support materials (Rerat et al., 2010). The trifluorotriazine reactions proceed analo-gously to the use of TsT in the methods presented in this section.

After the initial activation of a support with TsT, the most reactive chlorine of the two remaining can be blocked with the relatively weak nucleophilic amine on aniline to create a monofunctional derivative. The last remaining chlorine of the cyanuric chloride ring is the most stable to hydrolysis, but it is still able to effectively couple to nucleophilic groups on affinity ligands under relatively mild conditions. The monofunctional deriva-tive is best used for aqueous ligand coupling reactions with proteins or other biomolecules. However, ligands soluble in organic solvent may be coupled to the dichlo-rotriazinyl support (without blocking with aniline) and under nonaqueous conditions in the presence of an organic base to result in high-yield reactions devoid of any accompanying hydrolysis. Affinity ligands coupled

N N

NCl Cl

Cl

TsT(Trichloro-s-

triazine)

+


OH

O

NN

NCl

Cl

NH2

Aniline

O

NN

NCl

Nonaqueous

HN

Dichloro-s-triazine

derivative

Monochloro-s-triazine derivative

FIGURE 15.63 Activation of hydroxylic support with TsT and blocking with aniline to produce a mono-functional derivative.



using TsT activation result in more stable linkages rela-tive to leakage than those coupled using CNBr activa-tion (Hodgins et al., 1980).

TsT mediated coupling has been used to immobilize a variety of biomolecules and synthetic organic ligands, including the coupling of peptides onto polypropyl-ene membranes (Gérard et al., 2011), a general activa-tion method for the preparation of affinity membranes on polymers or carbohydrates (Avramescu et al., 2008), as an activating agent of amino-substituted agarose to create rationally designed affinity ligands (Roque and Lowe, 2008), and as an activator of aminopropyl silica for the preparation of immobilized affinity ligands (Luong and Scouten, 2008).

TsT Activation ProtocolThe following activation procedure should be car-

ried out in a fume hood with the appropriate personal protective equipment to prevent contact or inhalation of solvents or reagents.

1. Wash 100 ml of a hydroxyl-containing chromatography support material (such as crosslinked agarose) with water (at least several bed volumes) to remove preservatives and storage solution components. In a fume hood, sequentially wash the support into increasing concentrations of acetone-in-water (such as 25% (v/v), 50%, and 75% acetone/water) and finally with 100% dry acetone for at least 10 to 20 bed volumes to remove the last traces of water. Note: Dioxane or acetonitrile can be substituted for acetone depending on the compatibility of the matrix to a particular solvent.

2. Drain the support to a wet gel cake, resuspend it in an equal volume of solvent by mixing, and transfer the slurry to a 500-ml, three-necked, round-bottom flask placed in a heating mantle within the fume hood. Mix the slurry using an overhead stirring motor with a paddle stirrer and add to the flask a water-jacketed condenser connected to cold flowing water and a thermometer.

3. In the fume hood, stir the gel slurry and heat to 50°C using the heating mantle to slowly increase the temperature. Use care not to overheat, as acetone boils at 56 to 57°C.

4. After the gel slurry has equilibrated at 50°C, add 20 ml of 2-M N,N-diisopropylethylamine (DIEA) in acetone. N,N-dimethylaniline may also be used as the proton acceptor, but avoid using organic bases such as TEA, pyridine, N-ethyl morpholine, or lutidine, because these will precipitate with TsT in the solvent (Hodgins et al., 1980).

5. After 30 min of stirring, add 20 ml of 1-M TsT (highly purified) in acetone.

6. React for 1 h at 50°C with mixing.

7. In the fume hood, wash the activated support with acetone to thoroughly remove excess reactant and reaction byproducts. Wash with at least 10 to 20 bed volumes of solvent to ensure removal of the last traces of unreacted TsT and DIEA base.

8. Transfer the washed gel to a clean vessel and add 200 ml of 2-M aniline in acetone to block the most reactive chlorine on the triazine ring. Mix for 30 min at room temperature. Note: If the TsT-activated support is to be used to immobilize a solvent-miscible ligand, then it may be desirable not to block one of the two remaining acyl chlorides with aniline, because no hydrolysis will occur during the coupling reaction. If aniline blocking is not performed, then go to step 10.

9. Wash the TsT-activated support (now as the mono-chlorotriazine derivative) with at least 10 to 20 bed volumes of acetone to remove the last traces of unreacted aniline.

10. Store the activated support as a 50% slurry in dry acetone at 4°C until use.

TsT LIGAND-COUPLING PROTOCOLTsT-activated supports that are stored in acetone

or other anhydrous solvents may be used directly for coupling amine-containing ligands under nonaqueous conditions. In this case, a TsT-modified support that contains two remaining reactive chlorotriazine groups can be used (i.e., no blocking was done with aniline to eliminate the most reactive chlorine and leave only the least reactive one remaining). Alternatively, the activated gel may be filtered free of most solvent and reacted in aqueous buffer to couple proteins or other water soluble molecules. The following two protocols describe these methods in a general sense, but optimization should be performed to determine the best reaction conditions for a particular ligand and the optimal molar ratios for the best-performing affinity support. The reactions involved in coupling amine-containing ligands to TsT-activated supports are illustrated in Figure 15.64.

(A) Nonaqueous Coupling of Amine-Containing Ligands to TsT-Activated Supports1. In a fume hood, drain 100 ml of the TsT-activated

support of excess acetone (or other solvent) using a sintered glass filter funnel suspended in a suction filter flask. Do not allow the support to dry.

2. Dissolve the amine-containing ligand to be coupled in 100 ml of acetone (or another water-miscible solvent) at a concentration of 1 to 5 mg/ml, which is normally sufficient for the immobilization of a small organic compound. The optimal concentration of ligand to be used in the coupling reaction should be determined experimentally on small quantities of activated resin to identify the best affinity support



performance in its intended application. For organic solvent based reactions, add an organic base to the ligand solution, such as DIEA, to make a final concentration of 2-mM.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for at least 2 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. Longer reaction times may be done if appropriate to reach maximal yield of coupling.

4. Block unreacted active sites by the addition of ethanolamine (6.1 g) to the solution to make a 1-M solution. Mix for an additional 30 min.

5. Transfer the gel slurry to a sintered glass filter in the fume hood that is suspended in a suction filter flask and wash extensively (at least 10 bed volumes) with solvent to remove the remaining ligand and reaction byproducts. If the ligand is detectable, such as by using spectrophotometric methods, continue to wash the support until no further ligand is detected in the washings. Finally, drain the support of excess solvent by pulling a gentle vacuum on the filter flask while breaking up the support into small, finely divided pieces using a spatula, but be careful not to allow the matrix to dry out. Once the support is broken into small pieces, remove the vacuum and resuspend the gel in water with mixing. Continue to wash the

support with water until all traces of solvent have been removed. Additional washes with 1-M NaCl as well as low and high pH conditions may be carried out as appropriate to remove any noncovalently adsorbed ligand. Finally, wash with water and store the affinity support as a 50% slurry in water containing a preservative at 4°C until used.

(B) Aqueous Coupling of Biomolecules to TsT-Activated Supports1. Wash 100 ml of a TsT-activated support into water

and coupling buffer by suctioning off excess acetone storage solution and resuspending the gel into water. This can be done by pulling a gentle vacuum on the filter flask to filter off solvent while breaking up the support into small, finely divided pieces using a spatula, while being careful not to allow the matrix to dry out. Once the support is broken into small pieces, remove the vacuum and resuspend the gel in water with mixing. Continue to wash the support with water for at least 5 to 10 bed volumes. Finally, wash the support with 2 to 3 bed volumes of coupling buffer (0.1-M sodium borate, pH 8.5, containing 0.15-M NaCl).

2. Dissolve the protein or other amine-containing macromolecule to be immobilized in coupling buffer at a concentration of 1 to 20 mg/ml, or for

O

NN

NCl

Cl

O

NN

NCl

HN

Dichlorotriazinylderivative

Monochlorotriazinylderivative

+

+

O

NN

N


HN

NH


O

NN

N

HN

NH

Nonaqueous

Aqueous

H2N

H2N

Potential for two ligandsto couple through

secondary amine bonds

Single ligand couplingthrough secondary amine bond

FIGURE 15.64 Coupling an amine-containing ligand to a TsT-activated support.



small molecules, dissolve it at a concentration of 1 to 5 mg/ml. The optimal concentration of ligand that is used for the immobilization reaction should be determined experimentally to obtain the best performance of the affinity matrix after coupling. Avoid buffers or solution additives that contain amines, such as Tris, glycine, or imidazole, as these will compete with the ligand coupling reaction. Also avoid the presence of thiol-containing compounds such as DTT, because these will react with the chlorotriazine groups.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for at least 24 h at room temperature using an overhead paddle stirrer or by end-over-end rocking in a sealed container. The reaction may be performed at 4°C or at room temperature, depending on the stability of the ligand being immobilized; however, the coupling yield at 4°C is about 50% less than that at room temperature. For small-molecule reactions where the compound is thermally stable, the temperature may be increased to higher levels (i.e., as high as 45°C) to further increase the efficiency of the immobilization process.

4. Filter and wash the support with several bed volumes of water to remove most of the unreacted ligand and reaction byproducts. Transfer the support to a clean vessel and add 100 ml of 1-M ethanolamine, pH 9, with mixing to block the remaining active groups. Stir the reaction for 1 h at room temperature.

5. Filter and wash the affinity support to remove uncoupled ligand, blocking agent, and reaction byproducts using coupling buffer, water, 1-M NaCl, and again with water. Depending on the ligand being coupled, other wash solutions may be used to completely remove unreacted ligand, such as detergents, denaturants, and high or low pH conditions. Finally, store the affinity support in water containing a preservative at 4°C until use.

2.2. Thiol-Reactive Immobilization Methods

Although the majority of methods used to immo-bilize affinity ligands onto chromatography supports involve reactive groups that target amines, the use of site-directed chemistry that can target other functional groups such as thiols can have significant benefit in cer-tain instances (Domen et al., 1990). Thiols are typically present in proteins at more limited locations than amines and they can, therefore, be used to covalently link at selective sites within a macromolecule. Cystine disul-fides in proteins also can be mildly reduced to provide thiol groups for coupling even if a protein in its native state does not have an available free thiol, such as in the

reduction of disulfides within antibody hinge regions or in F(ab’)2 fragments (see Chapter 20). This strategy can result in the immobilization of proteins in areas away from binding sites or active centers, thus avoiding the blocking of these regions by being forced down toward the matrix during coupling instead of facing outward and available to interact with molecules in the mobile phase. Peptides can also be synthesized with a cysteine residue at one end of their amino acid sequences, thereby providing a functional handle to orient the immobilized peptide with the important binding end facing out from the support. Thiols can even be purposely added to pro-teins and molecules through the use of special modi-fication reagents to facilitate subsequent coupling to a thiol-reactive support (see Chapter 2, Section 4.1).

The many choices available in thiol-specific immobi-lization reactions described in this section can provide important options in designing an affinity support with the best possible performance for a particular applica-tion. Some of the methods form permanent or stable linkages with thiol-containing molecules by creating thioether bonds. Other methods are capable of forming reversible linkages using disulfide bonds, which subse-quently can be reduced to elute off the ligand from the support along with any interacting molecules. The fol-lowing sections describe the most common methods used for support activation and coupling to thiol-con-taining ligands for affinity chromatography. The reader is also directed toward the various other sections within this book that describe bioconjugate reagents for use with thiols to better understand the chemistry of thiol reactivity and coupling.

Iodoacetyl and Bromoacetyl ActivationHaloacetyl compounds have been used for decades

in the crosslinking, modification, and immobiliza-tion of thiol-containing molecules, especially for cova-lently linking to cysteine-containing proteins and peptides (Narayan et al., 2004; Wilhelmsen et al., 2004; Handlogten, et al., 2005; Kim and Hage, 2006a; Mallik et al., 2007). The reactivity of this group has its origin in the electron-withdrawing properties of the carbonyl oxygen of the carboxylate (note that the iodine atom of iodoacetyl compounds has an electronegativity approx-imately equivalent to that of the carbon to which it is attached). This effect causes an electrophilic center of partial positive charge on the carbon atom attached to the halogen. Nucleophiles such as thiols containing an unshared pair of electrons can attack this carbon result-ing in a potential nucleophilic substitution reaction, which causes displacement of the halogen with simulta-neous formation of a thioether bond with the thiol-con-taining compound.

Iodoacetyl groups in particular have been used to immobilize affinity ligands on all types of solid phases,



including chromatography supports, nonporous parti-cles, and planer surfaces for arrays (Camarero, 2006; Fu et al., 2011). Once a support is activated to contain iodo-acetyl groups it is relatively stable in aqueous solution, provided it is protected from light and reducing agents. Strong light exposure, especially sunlight, can cause the loss of the iodine atom (as HI) and rapidly eliminate coupling capacity. For this reason, activated supports should be stored in containers that prevent light trans-mittance to avoid this problem. Thiol-reducing agents can also react with the iodoacetyl groups and should be avoided, but even other reductants such as sodium cyanoborohydride, sodium borohydride, and phos-phine reducing agents (e.g., TCEP) can liberate HI and destroy activity, as well.

Bromoacetyl groups can also can be used to create thiol-reactive support materials. All of the reactions described in this section relative to iodoacetyl groups and their synthesis apply to bromoacetyl chemistry, too. To use bromo- instead of iodo-derivatives, substi-tute bromoacetate for use of iodoacetate in the prepara-tion of the activated support. The bromoacetyl group is more reactive than iodoacetyl, because the bromine atom is more electronegative than the iodine atom; however, in practice the two groups perform nearly identically in the immobilization of thiol-containing ligands onto chromatography supports. The iodoacetyl-activated supports are the most commonly cited in the literature for coupling thiol ligands.

Iodoacetyl groups can potentially react with any nucleophilic site in biomolecules or other ligands, including thiols, amines, and hydroxyl groups, depend-ing on the conditions of the reaction. The product of the reaction is an alkylation of the ligand by the C2 carbon atom of the acetyl group, which forms thioether, sec-ondary amine, or ether linkages with thiols, amines, or hydroxyls, respectively (Figure 15.65). Methionine thio-ether side chains are the most highly reactive toward alkylation and they can be modified even at acid pH conditions (pH 4.0) (Gundlach et al., 1959). Reaction of an iodoacetyl group with methionine yields an unsta-ble alkylation product, the carboxymethyl sulfonium salt of methionine, which subsequently can degrade by several routes into homocysteine, S-carboxymethyl homocysteine, or back to methionine (see Chapter 3, Section 2.1). Care should be taken using this method of coupling, therefore, if a methionine residue in a peptide or protein ligand represents a particularly important amino acid that must remain untouched by the immobi-lization process. Even if conditions are right to prevent amine or hydroxyl modification during the coupling of a thiol compound, the presence of methionine definitely may result in cross-reactions at this site.

Using iodo- or bromoacetyl-activated supports, thi-ols including cysteine will typically react under slightly

alkaline conditions that involve the use of a buffer in the range of pH 8.0 to 8.5. In this range, amine reactiv-ity will be very limited and hydroxyls will be virtually non-reactive. In fact, most protocols even make use of the amine-containing buffer Tris during the reaction, indicating that the specificity of the reaction toward cysteine thiols is very high. Increasing the alkalinity of the reaction can be used to purposely couple with amines (pH 10–12), while hydroxyls can be effectively targeted at pH > 12. Amine and hydroxyl group reactiv-ity may also be accompanied by the covalent modifica-tion of one of the imidazole ring nitrogens of histidine as well as the phenolic hydroxyl group of tyrosine residues. Thus, iodoacetyl supports potentially are one of the few immobilization chemistries that can have multi-purpose immobilization selectivity—the others being epoxy, TsT, and vinyl sulfone, which are able to couple with a variety of nucleophilic functional groups depending on the conditions used during the reaction. However, the targeting of amines or hydroxyls in the presence of thiols is not possible, because thiols have greater reactivity and will always be modified under higher pH conditions (unless the thiols are protected beforehand). The greatest benefit of using iodoacetyl activation is the potential to chemoselectively target

Iodoacetyl-activated support

S




Thioether linkage

Secondaryamine linkage

Ether linkage

HN

OI

HN

O

NH

HN

O

O

HN

O

Incr

easi

ng p

H

HS

H2N

HO

FIGURE 15.65 Reaction of an iodoacetyl-support with thiol-, amine-, and hydroxyl-containing supports.



only thiols (e.g., cysteines) at mild pH, thus leaving other functional groups alone.

Iodoacetyl supports are typically prepared from amine-containing matrices by coupling the carboxylate of iodoacetic acid to a terminal amine-containing spacer arm to form an amide bond. Thus, the initial step in the preparation of this support may be to modify an amine-reactive matrix with a diamine in large excess to form the terminal amino modifications suitable for reaction with iodoacetate. Many different diamine spacers can be used for this purpose, including diaminodipropylamine (DADPA), ethylene diamine (EDA; such as that used in the formation of MANAE supports), and the hydro-philic Jeffamine spacers containing short PEG groups (see the section on spacer arms, this chapter). The final coupling of the iodoacetate molecule onto the amino spacer can be performed using iodoacetic anhydride or NHS-iodoacetate, or through a carbodiimide (EDC) cou-pling procedure to attach the carboxylate to the amine through an amide bond. Mallik et al. (2007) prepared a thiol-reactive silica support using NHS–iodoacetate in the modification of an aminopropyl silane-coated silica particle (see Chapter 13, Section 2) to immobilize ligands for ultimate use in high-performance affinity chromatog-raphy (HPAC). The synthesis of a similar iodoacetate-agarose matrix made using the diamine spacer DADPA and attaching iodoacetate using EDC is described in the following protocol (Figure 15.66).

PREPARATION OF AN IODOACETYL-ACTIVATED SUPPORT

1. Prepare 100 ml of a DADPA-agarose support using periodate activation and diamine coupling through a reductive amination process as described elsewhere

in this chapter in the section on the preparation of spacer arm derivatives. Wash the support with water and then with several bed volumes of 0.1-M MES buffer, pH 4.7 (coupling buffer). Drain to a wet cake, but do not allow the gel to dry.

2. Dissolve 12 g of iodoacetic acid into 100 ml of coupling buffer (makes a 0.645-M solution) and adjust the pH back to 4.7 with base. Protect the compound and the solution from light to prevent degradation of the iodoacetate. Note: Bromoacetic acid may be substituted for iodoacetic acid by adding an equal mole amount (8.96 g) to the reaction slurry.

3. Add the wet gel cake to the iodoacetate solution with stirring.

4. With stirring, slowly add 10 g of EDC to the slurry to dissolve. React for 2 h at room temperature with constant mixing using an overhead paddle stirrer or end-over-end rotation in a sealed container. Protect the slurry from light by wrapping the vessel in aluminum foil.

5. Filter off the excess reaction solution and wash the activated gel with water, 1-M NaCl, and water (at least 10 bed volumes each) to remove unreacted compound and reaction byproducts. Store the iodoacetyl–agarose support as a 50% slurry in water containing a preservative at 4°C until use. Protect the gel from light to avoid decomposition.

LIGAND COUPLING TO IODOACETYL-ACTIVATED SUPPORTS

The first method described below is a generalized protocol that can be used to immobilize thiol-containing ligands onto iodoacetyl-activated supports. Molecules containing free thiols are notoriously susceptible to


O

H+ H2N N

HNH2

DADPA

NH

NH

NH2NaCNBH3

Amine-containing support

OI

O

Iodoacetate

NH

NH

NH


IO

EDC

FIGURE 15.66 Synthesis of iodoacetyl–DADPA–agarose using reductive amination to link the diamine spacer followed by EDC coupling of iodoacetate onto the terminal amine group.



oxidation in solution. Cysteine-containing peptides should be dissolved in nitrogen-purged and vacuum-degassed buffer solutions containing EDTA (at least 5 to 10-mM) to avoid oxygen-catalyzed or metal-catalyzed oxidation of the thiols to disulfides. This oxidation pro-cess can take place very quickly in solution and result in virtually no coupling of ligand to the support mate-rial. In addition, after the reduction of protein disul-fides to produce free thiols for coupling, the complete removal of the reducing agent is essential to prevent side reactions occurring with the iodoacetyl groups on the support.

The second procedure described below involves the conjugation of intact IgG or F(ab’)2 fragments to an iodoacetyl support after reduction of the disul-fide linkages between the heavy chains (Figure 15.67). The immobilization of antibodies or antibody frag-ments through thiols can result in coupling to sites that are away from the antigen binding areas, thus potentially preserving activity better than when using amine-reactive strategies. Using intact antibodies, the reduction process will result in a number of free thiols being produced, the amount of which is dependent on the concentration of the reducing agent and the type of antibody. Often, antibodies are most susceptible to reduction at the disulfides in the hinge region between the heavy chains, but reduction may also occur between

the heavy and light chains, which could result in dis-ruption of the antigen binding site. High concentrations of reducing agents used with antibodies could result in the complete dissociation of the heavy and light chain fragments, and if this preparation is used during the immobilization reaction with an iodoacetyl support the result might be little to no specific binding activ-ity after coupling. Therefore, controlling the amount of reductant added is important to the creation of thiols while not destroying the binding activity of the anti-body toward antigen. After reduction, the amount of thiols present within the ligand to be immobilized can be determined using Ellman’s reagent (see Chapter 2, Section 4.1).

(A) Coupling Thiol-Containing Ligands to Iodoacetyl-Activated Supports1. Wash 10 ml of an iodoacetyl-activated support with

water and then into coupling buffer using a sintered glass filter funnel suspended in a suction filter flask (coupling buffer: 50-mM Tris, 0.15-M NaCl, 10-mM EDTA, pH 8.5). The coupling buffer should be purged with nitrogen and degassed under vacuum to remove oxygen, which may oxidize the thiols and prevent coupling. Protect the activated resin and all solutions from light before and during the coupling reaction.

IgG Antibody

Reducingagent

Reduced antibody

+

Reduced antibody

Antibody immobilizedvia thioether bond

SO

pH 8.5

IO

HS

HS

SH

(A)

(B)


FIGURE 15.67 Reduction of antibody disulfides and coupling to an iodoacetyl-activated support.



2. Dissolve a thiol-containing ligand in 10 ml of coupling buffer at a concentration of at least 2 to 5 mg/ml. Determination of the optimal level of ligand concentration may be achieved experi-mentally by running a series of small reactions at different concentrations and determining which preparation performs best in the intended affinity chromatography application.

3. Add the ligand solution to the iodoacetyl-support and mix to thoroughly resuspend the resin in the solution. React for at least 1 h at room temperature (2–4 h at 4°C) with constant mixing.

4. Wash the gel with several bed volumes of coupling buffer to remove most of the excess uncoupled ligand. The amount of ligand not coupled may be determined by measuring the volume and concentration of the ligand in the pooled washes. This may be done using a protein assay or spectrophotometrically if the ligand has a characteristic spectral signature. The amount of ligand that has coupled to the matrix may then be determined by the difference. Drain the gel to a wet cake.

5. Block unreacted iodoacetyl active sites by adding to the gel 10 ml of a solution consisting of 50-mM cysteine (6.06 mg/ml) dissolved in coupling buffer. React for 30 min with stirring.

6. Wash the affinity resin thoroughly with water, 1-M NaCl, and again with water (at least 10 bed volumes each) to remove unreacted components. Store the support until use at 4°C as a 50% slurry in water containing a preservative.

(B) Coupling of Reduced IgG or F(ab’)2 Fragments to Iodoacetyl-Activated Supports1. Dissolve 1 to 10 mg of an IgG antibody or F(ab’)2

fragment in 1 ml of 0.1-M sodium phosphate, 0.15-M NaCl, 5-mM EDTA, pH 6.0.

2. Add a reducing agent [such as DTT, 2-mercaptoethylamine (2-MEA), 2-mercaptoethanol (2-ME), or tris(carboxyethyl)phosphine (TCEP)] to the antibody solution to give a final concentration of at least 5-mM. Mix to dissolve and incubate for 1.5 h at 37°C. Note: Higher concentrations of the reducing agent are sometimes used (i.e., up to 50-mM), but the higher the concentration the more likely will be the reduction of disulfides between the heavy and light chains, which may disrupt the three-dimensional structure of the antibody and destroy antigen binding capability. Optimization of the reductant concentration may have to be done to ensure the best performance of the resultant immunoaffinity support.

3. Remove excess reducing agent from the antibody solution by desalting using size exclusion

chromatography or by dialysis. During the chromatography or dialysis operation use coupling buffer consisting of 50-mM Tris, 0.15-M NaCl, 10-mM EDTA, pH 8.5, to equilibrate and elute protein through the gel filtration support or as the dialyzing solution. A gel filtration column should consist of at least 10 ml of a support having a molecular weight exclusion limit of no more than 5 to 10 kDa to ensure that the protein will come through in the void volume and be separated from the lower-molecular-weight reductant. A similarly sized dialysis membrane is appropriate. Complete removal of all reducing agent is essential to eliminate competition when coupling the reduced antibody to the iodoacetyl-activated support. Pool the fractions containing desalted protein peak from the gel filtration column or recover the dialyzed protein from the dialysis device. The solution may be concentrated if necessary using centrifugal concentrators to approximately 1 ml to maintain the desired concentration level of antibody in the coupling reaction.

4. Wash 1 ml of an iodoacetyl-activated support with water and then into coupling buffer by placing the resin into a drip column having a bottom porous frit and suspended in a test tube (coupling buffer: 50-mM Tris, 0.15-M NaCl, 10-mM EDTA, pH 8.5). If larger amounts of resin are used for the coupling reaction, then the use of a sintered glass filter funnel suspended in a suction filter flask is more appropriate for the washing steps. The coupling buffer should be purged with nitrogen and degassed under vacuum to remove oxygen, which may oxidize the thiols and prevent coupling. Protect the activated resin and all solutions from light before and during the coupling reaction.

5. Mix the washed iodoacetyl support with the reduced antibody solution and react in a sealed tube by gentle rotation for 1 h at room temperature. The tube should be wrapped in aluminum foil to prevent degradation of the iodoacetyl groups before the coupling reaction has occurred. If the antibody is sensitive to mixing, the tube can be rotated for the first 15 min and then allowed to incubate without continuous rotation. Every 5 min, gently resuspend the gel in the coupling solution to maintain a homogeneous slurry.

6. Wash the support with several bed volumes of coupling buffer to remove most of the not-coupled antibody. The washes may be analyzed versus the initial concentration of the antibody solution before coupling to determine the amount immobilized.

7. To block unreacted iodoacetyl groups, add to the washed resin 1 ml of 50-mM cysteine solution (6.06 mg/ml) prepared in coupling buffer. Mix and react for 30 min at room temperature.



8. Thoroughly wash the immunoaffinity support with coupling buffer, water, 1-M NaCl, and again with water to remove unreacted materials. Store the support until use at 4°C in water as a 50% slurry containing a preservative.

Maleimide ActivationThiol-containing affinity ligands can be immobilized

onto solid supports using maleimide-reactive groups. Maleimides long have been used in crosslinking and modification reagents to target thiols in cysteine-con-taining proteins for bioconjugation purposes (refer to the Index for many cross-references to this reactive group throughout this book). The specificity of the maleimide group toward thiols is excellent when reactions are per-formed at mildly basic pH. Side reactions might occur with amines if the pH is raised to a highly alkaline envi-ronment, but targeting thiols with maleimides is ele-gantly specific around physiological pH.

Maleimide groups can be formed on chromatogra-phy supports using an amine-containing spacer arm fol-lowed by reaction with a heterobifunctional crosslinker, which contains an amine-reactive NHS ester on one end and a maleimide group on the other end. Some exam-ples of these reagents include sulfo-SMCC (Chapter 6, Section 1.3), sulfo-GMBS (Chapter 6, Section 7), or NHS–PEGn–maleimide crosslinkers (Chapter 18, Section 1.2). An SMCC-based method was used to create maleimide groups on a silica support that had been modified with aminopropyl silane to form the requisite amino terminal spacers (Mallik et al., 2007). EMCS-based linkers have also been described to immobilize affin-ity ligands containing thiols (Kim and Hage, 2006b). However, a superior alternative for modifying sur-faces, particles, and chromatography supports with thiol-reactive maleimide groups is to use a PEG-based

crosslinker, which can add hydrophilicity to the spacer group formed on the matrix and thus limit nonspecific binding by avoiding aliphatic linkers. Building hydro-philic amine-terminal spacers off of a glycidol-modi-fied support (see section on periodate oxidation and reductive amination coupling, discussed previously) will also result in extremely biocompatible linkers, which can be used for modification with a maleimide heterobifunctional crosslinker. The reaction of NHS–PEG4–maleimide with a MANAE–agarose amine-con-taining spacer arm prepared from a glycidol-modified precursor is shown in Figure 15.68. Other amine-termi-nal spacer groups may also be used for this purpose to create a thiol-reactive support. Similar chemical cou-pling strategies have been used successfully for immo-bilizing affinity ligands onto surfaces (Houseman et al., 2003; Misra and Dwivedi, 2007).

The maleimide-reactive group undergoes rapid alkylation with a thiolate anion (–S−) in the pH range of 5.5 to 8.5 and displays second-order reaction kinetics (Tournier et al., 1998). Under these conditions, the major competing reaction is the potential of the maleimide ring to hydrolyze by opening to the maleamic acid derivative, which essentially inhibits effective cou-pling with thiols (see Chapter 19, Section 5). However, under standard coupling conditions the immobilization of a thiol ligand proceeds much faster than the rate of hydrolysis, yielding maximal density of ligand within 2 to 4 h at room temperature (Figure 15.69). The reaction of the blocked amino acid N-acetylcysteine with the maleimide group of sulfo-SMCC was found to be 50% complete within 20 min at pH 6.5 and totally complete within 10 min at pH 8.5 (Tournier et al., 1998). Allowing reactions to go for a longer time for immobilization reactions onto porous chromatography supports is good practice, as the diffusion of large protein ligands

O

HN

NH2


primary amines

+

OO

ON

O

O

OO

ON

O

O

NHS-PEG4-maleimide

O

HN

NH

O

O

O

O

ON

O

O

Maleimide-activatedsupport

FIGURE 15.68 Preparation of maleimide-agarose by the reaction of NHS–PEG4–maleimide with MANAE–agarose.



into the particles will increase the reaction times neces-sary to obtain optimal yields.

A survey of the literature on maleimide immobiliza-tion of thiol-containing affinity ligands indicates that this technique has mainly been used to activate and couple molecules to surfaces or nonporous particles (Hwang et al., 2011; Cellet et al., 2012). This is not a limi-tation of the maleimide group with respect to its util-ity, but a realization that the cost of materials to form a maleimide-reactive chromatography support may be prohibitive for large-scale use. The creation of other thiol-reactive supports, such as iodoacetyl, discussed previously, involves the use of reagents that are avail-able in multi-gram quantities at relatively inexpensive price points. By contrast, maleimide crosslinkers used to form the support derivatives described in this sec-tion are much more expensive on a per-gram basis. Therefore, if small amounts of resin are being acti-vated, this method is eminently appropriate; however, if quantities > 10 ml of support are to be prepared, then it might be better to consider another coupling method due to the price of the maleimide crosslinker.

PREPARATION OF A MALEIMIDE-ACTIVATED SUPPORT

The following protocol requires the use of an amine-containing chromatography support, which may be prepared by the methods discussed in the section on spacer arms elsewhere in this chapter.

Protocol1. Wash 10 ml of an amine-containing support material

(such as MANAE–agarose) with water to remove storage solutions and then with several bed volumes of 0.1-M sodium phosphate, pH 7.2 (activation buffer). Finally, suspend the support in an equal volume of activation buffer with mixing.

2. While mixing the suspended amine-containing gel, add 218 mg sulfo-SMCC (Thermo Scientific Pierce)

(equivalent to about 50 μmoles/ml gel). Alternatively, an equivalent mole quantity of NHS–PEG4–maleimide may be added instead of sulfo-SMCC to create a more hydrophilic support environment due to the presence of the PEG-based spacer arm.

3. React with constant mixing for 1 h at room temperature. The mixing may be done using an overhead stirring paddle or end-over-end rotation in a sealed container.

4. Quickly wash the activated support with several bed volumes of activation buffer, 1-M NaCl, and with water to remove unreacted crosslinker and reaction byproducts. Drain to a moist cake. To prevent hydrolysis of the maleimide groups, use the activated support immediately to immobilize a thiol-containing ligand according to the following protocol.

LIGAND COUPLING TO MALEIMIDE-ACTIVATED SUPPORTS

1. Prepare 10 ml of a ligand solution for immobilization by dissolving in coupling buffer (0.1-M sodium phosphate, 0.15-M NaCl, 10-mM EDTA, pH 7.2) a protein containing one or more free cysteine thiols or a small molecule ligand having an available thiol at a concentration of 1 to 20 mg/ml for the protein or 2 to 5 mg/ml for the small ligand. Alternatively, the small molecule may be reacted with the support at a concentration of 50 to 100 μmoles/ml gel. The thiols on proteins may be generated by the use of a reducing agent to cleave disulfides (see the protocol described previously for coupling reduced antibodies to iodoacetyl-activated supports) or through the use of a thiolation modification reagent (see Chapter 2, Section 4.1).

2. Add 10 ml of the washed, maleimide-activated support prepared above to the ligand solution with stirring to resuspend the matrix. React at room temperature for at least 2 h with constant mixing using an overhead paddle stirrer or end-over-end mixing in a sealed container.

3. Wash the support with several bed volumes of coupling buffer to remove most of the not coupled ligand. Drain to a moist cake. The washes may be analyzed versus the initial concentration of the ligand in the coupling solution before the reaction was initiated to determine the amount immobilized.

4. To block unreacted maleimide groups, add to the washed resin 10 ml of 50-mM cysteine solution (6.06 mg/ml) prepared in coupling buffer. Mix and react for an additional 30 min at room temperature.

5. Thoroughly wash the affinity support with coupling buffer, water, 1-M NaCl, and again with water to remove unreacted materials. Store the support until use at 4°C in water as a 50% slurry containing a preservative.

Maleimide-activatedsupport N

O

O+


S

Ligand immobilizedthrough thioether

linkage

N

O

O

HS

FIGURE 15.69 Coupling of thiol-containing ligands to a maleimide-activated support.



Divinyl Sulfone ActivationDivinyl sulfone (DVS) is a highly reactive homo-

bifunctional compound that can be used to activate hydroxylic matrices for coupling to a wide variety of affinity ligands (Porath, 1974). It has also been used as a crosslinking agent to increase the physical strength of hydroxylic supports and as a gelling agent to form large-scale crosslinks in polymers for industrial appli-cations (Kriegel, 2006). Activation of an agarose sup-port with DVS in large excess occurs under alkaline pH conditions with modification of the hydroxyls present on the support. This process results in the reaction of one side of the DVS molecule with an avail-able hydroxyl group on the support, which proceeds through a Michael-type addition to the double bond. At the same time, the formation of reactive vinyl sul-fone groups occurs due to the presence of the unreacted end of the molecules remaining free (Figure 15.70). If the reaction is performed with a large excess of DVS, it will occur with minimal crosslinking within the matrix, although closely spaced hydroxyl groups may indeed link together and increase the overall structural rigidity within a resin at the same time as activation occurs. The resultant vinyl sulfone reactive groups are stable for storage in aqueous solution as long as no nucleophiles are present in the storage solution. This simple activa-tion process combined with the excellent hydrolytic stability of DVS-activated supports (at 4°C) makes this method of ligand coupling highly attractive, especially for immobilizing ligands containing thiols or hydroxyls.

Supports activated to contain vinyl sulfone groups can be used to immobilize ligands containing nucleo-philic thiol, amino, or hydroxyl groups with good link-age stability (Lihme et al., 1986). Ubrich et al. (1992) also found that DVS-mediated coupling resulted in one of the most stable affinity supports for the immobilization of antibodies. Highly activated supports typically can couple proteins at densities of up to 30 to 50 mg/ml gel,

depending on the quantity of protein initially reacted. Small-molecule ligands can also be coupled at high mole densities. Thiol-containing ligands are the most reac-tive and are able to couple with the vinyl sulfone groups at a pH of 6 to 8, while forming thioether linkages of excellent stability. Amine-containing ligands may also be immobilized onto vinyl sulfone supports within the somewhat higher pH range of 8 to 10, which results in a secondary amine bond upon addition to the double bond. Finally, hydroxyl groups on ligands such as those on polysaccharides, carbohydrates, and glycans may be linked to vinyl sulfone supports at pH values > 10 to create ether bonds (Figure 15.71). The pH modulation of vinyl sulfone to react with a single nucleophile type is only limited by the presence of other more nucleophilic groups. In other words, an amine cannot be targeted in the presence of a thiol without coupling also taking place at the thiol site. Similarly, hydroxyl functionalities cannot be targeted for coupling in the presence of other amines or thiols, because the more nucleophilic groups will react preferentially or in addition to the hydroxyl.

The yield of protein coupling to a vinyl sulfone-activated matrix may be enhanced by the addition of relatively high concentrations of PEG added to the reac-tion medium (5–7% PEG for antibodies, 7–10% PEG for most other proteins; Mini-Leak protocol, Kem-en-tec). In addition, similar enhanced coupling rates and

Hydroxyl-containingsupport

+

Divinyl sulfone

Vinyl sulfone-activated support

S

O

O

OS

O

O

OH-

OH

FIGURE 15.70 Activation of a hydroxylic support with divinyl sulfone.




Thioether linkage


Ether linkage

HN

O

Incr

easi

ng p

H


OS

O

O

SOS

O

O

OS

O

O

OS

O

O

HS

H2N

HO

FIGURE 15.71 Reaction of vinyl sulfone-activated supports with thiol-, amine-, and hydroxyl-containing ligands.



yields can be observed with the addition of a lyotropic salt (sodium sulfate, potassium sulfate, ammonium sul-fate, or potassium phosphate, > 0.5-M). Both the PEG and high salt concentrations have the effect of push-ing proteins out of solution and toward the surfaces of the activated support where they can more effectively react with the vinyl sulfone groups. This effect is also observed with azlactone-activated supports and with epoxide coupling reactions, described elsewhere in this chapter. Some optimization of the level of PEG or salt addition should be carried out to obtain the best rate of reaction for a given ligand to be immobilized.

The degree of vinyl sulfone-activated species within a chromatography support may be determined by reac-tion of a small sample of the gel with a large excess of sodium thiosulfate (Porath et al., 1975; Lihme and Boenisch, 1996; Subramanian, 2000). The thiosulfate anion reacts with the vinyl groups and generates OH− ions (Figure 15.72). Subsequent titration with HCl of the amount of hydroxide ions released provides a measure-ment of the quantity of vinyl sulfone groups originally present within the matrix.

Divinyl sulfone-activated supports are more reac-tive than epoxide-activated supports, but the abilities of both immobilization methods are similar in that they can be made to couple with thiols, amines, or hydrox-yls simply by modulating the pH of the reaction. The resultant sulfone bridge created between the support and the ligand is stable under physiological conditions, but it can be purposely cleaved by exposure to highly alkaline conditions (see Chapter 9, Section 5). Retro-Michael cleavage of base labile sulfones can be accom-plished by mixing the support as a 50% slurry in 0.1-M sodium carbonate (or sodium phosphate), pH 11.6, and heating the stirred gel suspension at 37°C for at least 2 h (Zarling et al., 1980). Longer incubation periods may be required for complete reversal of immobilized ligands. The reversible nature of the support indicates that DVS-coupled affinity ligands should not be exposed to con-ditions exceeding pH 8.5 for long periods of time to avoid potential ligand leakage through cleavage of the sulfone linker arm. However, under normal use and storage conditions the sulfone linker arm is very stable and provides a robust affinity support suitable for long term use.

DVS-activated supports have often been used to immobilize a wide variety of biological molecules to solid supports (Pepper, 1994; Morales-Sanfrutos et al., 2010), including to covalently link glycans onto surfaces (Cheng et al., 2011), to activate biodegradable polymers for coupling bioactive molecules (Wang et al., 2011), in the activation of silica particles for the coupling of thiol-containing affinity ligands (Ortega-Munoz, 2010), and in the preparation of reactive polymers (Lihme and Boenisch, 1996).

DVS activation is also critical in the preparation of thiophilic adsorbents that are used in the isolation of immunoglobulins. The principle of thiophilic inter-action chromatography involves an affinity for cer-tain regions within the Fc fragment of antibodies by a ligand consisting of a sulfone in proximity to a thio-ether group, with or without the addition of an aro-matic group nearby in the structure (Porath et al., 1985; Hutchens and Porath, 1986, 1987a,b; Belew et al., 1987; Porath, 1987; Porath and Belew, 1987; Nopper et al., 1989; Lihme and Heegaard, 1991; Knudsen et al., 1992; Hardouin et al., 2007). The interaction was discovered entirely by accident, as the affinity does not involve a biospecific interaction, but a synthetic ligand which likely binds through hydrophobic contacts with pock-ets within the Fc domains of immunoglobulins. A thio-philic affinity resin can be prepared simply by coupling 2-mercaptoethanol to a DVS-activated support accord-ing to the following protocols (Figure 15.73). The resul-tant support can be used to capture antibodies in the presence of high concentrations of a lyotropic salt (at least 0.5-M potassium sulfate), which serves to enhance hydrophobic interactions. Subsequent elution of bound antibody is carried out merely by elimination of the salt from the buffer, thus making this method extremely mild for purifying IgG antibodies. The capacity of this affinity support for human IgG can be as high as 20 mg/ml of thiophilic gel.

PREPARATION OF A DVS-ACTIVATED SUPPORT

The activation protocol described below should be done with caution, as divinyl sulfone is a highly reac-tive and toxic compound. All operations should be carried out in a fume hood and with the use of proper personal protective equipment.


OS

O

O

Sodiumthiosulfate(Na2S2O3) O

S

O

O

+ OH-SS

O

O

O

Thiosulfate derivativewith OH- released

FIGURE 15.72 Reaction of sodium thiosulfate with vinyl sulfone to generate hydroxyl ions.



Protocol1. Wash 100 ml of a support material containing

hydroxyl groups (such as agarose) with water (at least 5 bed volumes) to remove storage solutions and preservatives. Typically, a non-crosslinked agarose resin can be used for the activation, because crosslinking will occur during the activation step and create a more rigid matrix. However, other natural or synthetic polymeric, hydroxyl-containing supports or inorganic supports containing hydroxyl groups may be used as well, whether crosslinked or not. Use a sintered glass filter funnel suspended in a suction filter flask while pulling a gentle vacuum to facilitate the washing steps. After washing the gel, suction to a moist cake by removing excess water, but do not allow the support to dry.

2. Suspend the gel in 100 ml of 0.5-M sodium carbonate (no pH adjustment) and mix in a fume hood using an overhead paddle stirrer.

3. Add 10 ml of divinyl sulfone dropwise to the stirring gel slurry over a period of about 15 min.

4. Continue to react for 1 h at room temperature with constant mixing.

5. Wash the activated gel extensively with water to remove excess DVS (at least 20 bed volumes). Storage of the activated support may be achieved as a 50% slurry in water containing a preservative (e.g., 1,1,1-trichloro-2-methyl-2-propanol) at 4°C. Properly stored, the support should remain stable up to 2 years with minimal loss of vinyl sulfone coupling activity.

LIGAND COUPLING TO DVS-ACTIVATED SUPPORTS

The following ligand-coupling procedures describe the use of a DVS-activated support (10 ml) for the coupling of a thiol-, amine-, or hydroxyl-containing ligand. This may include the immobilization of proteins, carbohydrates, or other molecules that contain these functional groups. The immobilization through thiols may be performed in the presence of other amines or hydroxyl groups on the ligand; however, coupling through amines must be done

while avoiding the presence of thiols, as they will pref-erentially react at a faster rate than amines. Similarly, the coupling of hydroxyls should be done while avoiding the presence of thiols or amines on the ligand.

(A) Coupling of Thiol-Containing Proteins or Ligands1. Prepare 10 ml of a thiol-containing ligand solution

by dissolving in coupling buffer (0.1-M sodium phosphate, 0.15-M NaCl, 10-mM EDTA, pH 7.5) a protein containing one or more free cysteine thiols or a small molecule ligand having an available thiol at a concentration of 1 to 20 mg/ml for the protein or 2 to 5 mg/ml for the small ligand. Notes: The EDTA chelator is present to inhibit metal-catalyzed oxidation of the thiols to disulfides. Alternatively, a small molecule may be reacted with the support at a concentration of 50 to 100 μmoles/ml gel. The thiols on proteins may be generated by the use of a reducing agent to cleave disulfides (see the protocol described previously for coupling reduced antibodies to iodoacetyl-activated supports) or through the use of a thiolation modification reagent (see Chapter 2, Section 4.1). Complete removal of any thiol-containing reducing agents must be performed before attempting to immobilize the reduced protein to avoid competition during the coupling reaction. The coupling buffer may also be formulated to contain a lyotropic salt, such as sodium sulfate, at a concentration of at least 0.5-M to increase the rate of reaction when immobilizing proteins. Alternatively, the addition of PEG (MW 20 kDa) to the coupling buffer may be used for this purpose at a concentration of 5 to 7% for antibodies or 7 to 10% for most other proteins.

2. Wash 10 ml of the DVS-activated support prepared above with water to remove any preservatives from the storage solution and then wash with coupling buffer (at least 2 bed volumes). Drain to a wet cake.

3. Add the gel to the ligand solution with stirring (total slurry volume: 20 ml). React overnight at room


OS

O

O

OS

O

O

SOH

HSOH

2-Mercaptoethanol

+Thiophilic support containinga thioether next to a sulfone

FIGURE 15.73 Preparation of a thiophilic adsorbent through coupling of 2-mercaptoethanol to DVS-activated supports.



temperature or for at least 18 h with constant mixing using an overhead paddle stirrer or end-over-end mixing in a sealed container. The reaction may also be performed at 4°C for sensitive molecules or proteins, but the reaction time may have to be extended to obtain the same levels of coupling.

4. Wash the support with several bed volumes of coupling buffer (without the lyotropic salt or PEG present) to remove most of the not-coupled ligand. The washes may be analyzed versus the initial concentration of ligand in the coupling solution before the reaction was started to determine the amount that was immobilized. Continue to wash with several bed volumes of water and drain to a wet cake.

5. To block unreacted vinyl sulfone groups, add the washed resin to 10 ml of 0.1-M cysteine solution (12.12 mg/ml) prepared in 0.1-M sodium bicarbonate, pH 8.6. Alternatively, use a blocking solution consisting of 0.2-M ethanolamine prepared in the same buffer. Mix and react for an additional 2 h at room temperature. Note: Do not block the support with 2-mercaptoethanol, because coupling this molecule may result in the creation of a number of thiophilic interaction sites within the matrix, which could cause nonspecific binding with antibodies and other molecules, depending on the conditions of the affinity chromatography being done.

6. Thoroughly wash the affinity support with 0.1-M sodium phosphate, pH 7.5, water, 1-M NaCl, and again with water to remove unreacted materials. Store the support until use at 4°C in water as a 50% slurry containing a preservative.

(B) Coupling of Amine-Containing Proteins or Ligands1. Prepare 10 ml of a amine-containing ligand solution

by dissolving in coupling buffer (0.1-M sodium bicarbonate, pH 8.6) a protein containing one or more free amines or a small molecule ligand having an available amine at a concentration of 1 to 20 mg/ml for the protein or 2 to 5 mg/ml for the small ligand. Alternatively, the small molecule may be reacted with the support at a concentration of 50 to 100 μmoles/ml gel. Note: The coupling buffer may also be made up to contain a lyotropic salt, such as sodium sulfate, at a concentration of at least 0.5-M to increase the rate of reaction when immobilizing proteins. Alternatively, the addition of PEG (MW 20 kDa) to the coupling buffer may be used for this purpose at a concentration of 5 to 7% for antibodies or 7 to 10% for most other proteins.

2. Wash 10 ml of the DVS-activated support prepared as described previously with water to remove any preservatives from the storage solution and then

wash with coupling buffer (at least 2 bed volumes). Drain to a wet cake.

3. Add the gel to the ligand solution with stirring to resuspend the matrix (total slurry volume: 20 ml). React overnight (for at least 18 h) at room temperature or at 4°C with constant mixing using an overhead paddle stirrer or end-over-end mixing in a sealed container.

4. Wash the support with several bed volumes of coupling buffer (without the lyotropic salt or PEG present) to remove most of the not-coupled ligand. The washes may be analyzed versus the initial concentration of ligand in the coupling solution to determine the amount immobilized. Continue to wash with several bed volumes of water and drain to a wet cake.

5. To block unreacted vinyl sulfone groups, add the washed resin cake to 10 ml of 0.2-M ethanolamine solution prepared in 0.1-M sodium bicarbonate, pH 8.6. Note: After the addition of ethanolamine to the buffer, adjust the pH back to 8.6 with 6-N HCl solution while maintaining the temperature near ambient using an ice bath if necessary. Mix and react for an additional 2 h at room temperature.

6. Thoroughly wash the affinity support with 0.1-M sodium bicarbonate, water, 1-M NaCl, and again with water to remove unreacted materials. Store the support until use at 4°C in water as a 50% slurry containing a preservative.

(C) Coupling of Carbohydrates or Hydroxyl-Containing Ligands1. Prepare 10 ml of a hydroxyl-containing ligand

solution (such as a carbohydrate, polysaccharide, or glycan) by dissolving it in coupling buffer (0.1-M sodium carbonate, pH 11.0) at a concentration of up to 100 mg/ml for mono- or disaccharides, or at least 1 to 20 mg/ml for larger polysaccharides or glycans. Alternatively, a hydroxyl-containing ligand may be reacted with the support at a concentration of 50 to 100 μmoles/ml gel. Notes: The more concentrated the ligand solution, the better will be the coupling yield of the reaction and ultimate density of ligand on the support. However, for the immobilization of carbohydrates to be used in the affinity capture of lectins or other carbohydrate binding proteins, it may be optimal to immobilize the ligand at lower than maximal densities. This is due to the fact that many lectins contain more than one binding site for a carbohydrate, and multi-site interactions on the affinity support will have a tendency to create strong avidity, which may be difficult to reverse for the efficient elution of interacting proteins. Thus, some degree of experimental optimization of ligand density may have to be performed to obtain the best



performance of the affinity support in its intended application.

2. Wash 10 ml of the DVS-activated support prepared previously with water to remove any preservatives from the storage solution and then wash with coupling buffer (at least 2 bed volumes). Drain to a wet cake.

3. Add the gel to the ligand solution with stirring (total slurry volume: 20 ml). React overnight at room temperature (for at least 18 h) or at 4°C with constant mixing using an overhead paddle stirrer or end-over-end mixing in a sealed container. Reactions done at 4°C may take extended time periods to reach the same level of coupling yield as those done at room temperature.

4. Wash the support with several bed volumes of coupling buffer to remove most of the not coupled ligand. The washes may be pooled and analyzed versus the initial concentration of ligand in the coupling solution before the reaction was initialized to determine the amount of ligand immobilized. Drain the gel to a wet cake.

5. To block unreacted vinyl sulfone groups, add the washed resin cake to 10 ml of 0.2-M ethanolamine solution prepared in 0.1-M sodium carbonate, pH 11.0. (Note: After the addition of ethanolamine to the buffer, adjust the pH back to 11.0 with 6-N HCl, if necessary. Mix and react for an additional 2 h at room temperature.

6. Thoroughly wash the affinity support with 0.1-M sodium carbonate, water, 1-M NaCl, and again with water to remove unreacted materials. Store the support until use at 4°C in water as a 50% slurry containing a preservative.

(D) Coupling of 2-Mercaptoethanol to Make a Thiophilic Resin

The following protocol represents one of several methods available to form a thiophilic affinity support for the purification of immunoglobulins and other pro-teins able to interact with the ligand.

1. Wash 10 ml of the DVS-activated support prepared previously with water to remove any preservatives from the storage solution and then wash with at least 2 bed volumes of coupling buffer (0.1-M sodium carbonate, pH 9.0). Drain to a wet cake.

2. In a well-ventilated fume hood, suspend the washed activated support in 9 ml of coupling buffer and add 1 ml of 2-mercaptoethanol (stench!) with stirring.

3. React overnight (at least 18 h) at room temperature with constant mixing. For small volumes of gel, mixing may be accomplished by end-over-end rotation in a sealed container. For larger volumes of gel the use of an overhead paddle stirrer is

appropriate; however, due to the unpleasant odor of 2-mercaptoethanol a sealed round-bottom flask should be used for the gel slurry during mixing operations.

4. Wash the resin thoroughly in the fume hood with coupling buffer, 1-M NaCl, and water. Continue the washing until no odor is detectable from 2-mercaptoethanol. Store the thiophilic support until use at 4°C in water as a 50% slurry containing a preservative.

Pyridyl Disulfide ActivationPyridyl disulfide groups can react with thiol-contain-

ing molecules by disulfide exchange to form new mixed disulfide linkages. This group long has been used as a thiol-reactive end in crosslinking reagents and protein modification agents, such as the popular heterobifunc-tional compound SPDP (Chapter 6, Section 1.1). Pyridyl disulfide groups also can be formed on chromatogra-phy supports to immobilize thiol-containing ligands for affinity separations through a simple reaction process (Cuatrecasas, 1970; Egorov et al., 1975; Carlsson et al., 1976; Ngo, 1989). The similar reagents 4,4′-dipyridyl disulfide (Grassetti and Murray, 1967) or 2,2′-dipyridyl disulfide (Brocklehurst et al., 1973) can be used as acti-vation agents to form reactive pyridyl disulfide groups on resins. Reaction of these compounds in excess with a thiol-containing support material results in the forma-tion of pyridyl disulfide groups and activation of the support for disulfide exchange reactions with a thiol-containing ligand (Figure 15.74). The requirement for activation using these reagents is to first form free thiol groups on a support using spacer arms that terminate in thiol groups. One method of creating such a thiol-containing matrix is to couple the bis-thiol compound DTT to an epoxy-activated support (see the section Epoxide Activation, this chapter). Another potential approach to forming terminal thiol groups is to modify an amine-containing support with a thiolation reagent

Thiol-containingsupport

2,2'-Dithiodipyridine(or 2,2'-Pyridyl disulfide)

+

Pyridyldithiol-activated support

SH

NS

SN

SS N

FIGURE 15.74 Activation of a thiol-containing support with 2,2′-pyridyl disulfide.



such as SATA, Traut’s reagent, or homocysteine thio-lactone (Chapter 2, Section 4.1), which form free or pro-tected thiols upon reaction with the amines.

Amine-containing supports may also be modified with heterobifunctional crosslinking agents to form the pyridyl disulfide reactive groups directly from an amine-terminal spacer (Brogan and Schoenfisch, 2005). In this case, reaction of the terminal amines with the crosslinking reagent SPDP (Chapter 6, Section 1.1) through its NHS ester end creates amide bond linkages with the amines on the support, which then forms short spacer arms that terminate in pyridyl disulfide groups. An even better option is to use an NHS–PEGn–pyridyl disulfide crosslinker, which has a hydrophilic spacer arm in its construction to create a matrix having low nonspecific binding characteristics (Chapter 18, Section 1.2) (Figure 15.75). This activation strategy may be the most straightforward, but it also is likely the most expensive due to the potential cost of the crosslinking reagent. For making small amounts of support material (i.e., <10 ml) the use of a pyridyl disulfide-containing

crosslinker is cost effective and a simple route to pro-duction of the activated support. It also works well for activation of surfaces or small nonporous particles. For making larger quantities of resin materials, however, it may be best to use the methods that employ the dipyri-dyl disulfide activation reagents, because these com-pounds are much less expensive.

Perhaps the simplest method of introducing pyridyl disulfide groups into a chromatography support is to use the reagent PDEA, which is 2-(2-pyridinyldithio)ethaneamine. This small compound has an amine on one end and the thiol-reactive group on the other end. An amine-reactive chromatography support may be used to couple this reagent in high yield, thus forming the pyridyldithiol groups in a single step. Figure 15.76 illustrates the coupling of PDEA to a succinimidyl-carbonate-activated support, which reacts with amines within the pH range of 7 to 9 to give carbamate link-ages (see the associated section, this chapter). Renberg et al. (2005) used PDEA to modify a sensor chip for the immobilization of thiol-containing Affibodies, which

Amine-containingsupport

NHS–PEG4–pyridyldithiolcrosslinker

+


NH2

HN

NO

O

O

O

OO

O S

OS NO

HN

O

OO

O S

OS NO

HN

FIGURE 15.75 Use of NHS–PEGn–pyridyl disulfide to create pyridyl disulfide groups from an amine-containing support.

SS NH2N

PDEA;2-(2-Pyridinyldithio) ethaneamine

Succinimidyl carbonate-activated support

+

O O

ON

O

O

OHN

OS

S N


FIGURE 15.76 Use of PDEA to form pyridyl disulfide groups on an amine-reactive support.



are synthetic peptides having antibody-like binding properties toward antigens. The use of PDEA is also a recommended method of coupling thiol affinity ligands to a Biacore sensor chip, because the chip can be regen-erated after use. PDEA-modified agarose supports have also been used to immobilize enzymes for use a bio-reactors (Mansfeld and Ulbrich-Hofmann, 2000). The amino group of PDEA was coupled to CNBr-activated agarose or through an aminocaproic acid spacer using the EDC/NHS coupling reaction to yield the final thiol-reactive derivative (Chapter 4, Section 1).

Once pyridyl disulfide groups are formed on a sup-port, they are stable for long-term storage in aqueous solution provided no reducing or oxidizing agents are present. The reaction of these groups with thiol-contain-ing affinity ligands results in the formation of reversible disulfide bonds with release of the chromogenic leaving group pyridine-2-thione (Figure 15.77). Once the leav-ing group is displaced by a thiol ligand, it cannot go back and again react with the disulfide groups formed on the matrix, because the thione double bond makes the sulfur unreactive toward disulfide exchange. The release of this group also can be used to estimate the degree of coupling by measurement of its absorbance at 343 nm, provided the ligand does not absorb in this region. The reaction of a thiol-containing ligand with the pyridyl disulfide groups on the support will occur

within the pH range of 4 to 8, with optimal reaction kinetics observed at pH 7 to 8 (Carlsson et al., 1978).

Pyridyl disulfide immobilization is an alternative to the permanent coupling methods discussed previ-ously for thiol ligands in that a coupled molecule can be subsequently removed from the support by treat-ment with a disulfide reducing agent. Disulfide immo-bilized ligands may be cleaved from the support by incubation with 25- to 50-mM DTT for 2 h at room tem-perature. This feature builds the potential for reversible covalent capture and release of ligand-target molecule interacting pairs under mild chromatography condi-tions. Therefore, this may be a valuable technique for co-immunoprecipitation (co-IP) assays in the study of protein interactions, because it allows for the isolation of these proteins under non-denaturing conditions. After a bound ligand is released, the support once again contains free thiol groups that then can be re-activated to form pyridyl disulfide groups for another immobi-lization reaction, thus allowing complete reusability of the support.

Pyridyl disulfide immobilization has also been used to couple thiol-containing antibodies, antibody frag-ments, proteins, and other molecules to polymers and solid phases of all types. For instance, Iwata et al. (2007) used pyridyl disulfide groups on polymer brushes to immobilize Fab’ fragments containing thiol groups

OHN

OS

S N



+O

HN

OS

Ligand immobilized throughdisulfide bond

S

HN S

Pyridine-2-thione

DTT

OHN

OSH

Ligand released by reductionof the disulfide bond

HS+

A343nm

HS

FIGURE 15.77 Coupling a thiol-containing ligand to a pyridyl disulfide-activated support with subsequent removal by treatment with a reducing agent.



within their hinge region. Vázquez-Dorbatt et al. (2009) formed these active groups within a glycopolymer by reaction of a pyridyl disulfide-containing initiator using atom transfer radical polymerization (ATRP) to cre-ate a copolymer matrix that could immobilize double-stranded siRNA, which had been previously modified to contain a terminal thiol group. Metal surfaces have also been activated with pyridyldithiol groups to allow affin-ity ligands to be coupled for molecular recognition force microscopy measurements (Brogan and Schoenfisch, 2005). In addition, a pyridyl disulfide-activated support can be used for the proteomics study of cysteine-contain-ing peptides by reversible capture and elution with sub-sequent analysis by mass spec (Shen et al., 2003).

PREPARATION OF A PYRIDYL DISULFIDE-ACTIVATED SUPPORT

The following methods for the preparation of a pyri-dyldithiol support represent different options for build-ing this reactive group onto various base supports. The same general methods may be used to activate particles or surfaces with excellent results. The level of reactive pyridyl disulfide groups formed on a support may be controlled by adjusting the amount of functional groups initially created on the support, which are subsequently modified to create the reactive groups. The following methods use either amine- or thiol-terminal spacer arms on a support to form the final reactive pyridyl disulfides.

(A) Activation using 2,2′-Dipyridyl DisulfideThis protocol will work well using either 2,2′-dipyri-

dyl disulfide or 4,4′-dipyridyl disulfide. These com-pounds may be obtained commercially (Acros, Aldrich) or synthesized from the corresponding mercaptopyri-dine precursor according to the method presented in Hermanson et al. (1992).

1. Prepare 10 ml of a support containing a thiol-terminal spacer using the methods described within the section on spacer arms in this chapter. The support should contain 10 to 50 μmoles –SH groups/ml of gel. Wash the support with water to remove storage solution components, if necessary, and drain to a wet cake.

2. In the fume hood, prepare 10 ml of an acetone:water (1:1, v/v) solution containing 1 g of 2,2′-dipyridyl disulfide dissolved into it.

3. Add the gel to the 2,2′-dipyridyl disulfide solution with stirring and react at room temperature for 30 min with constant mixing in the fume hood. For small amounts of gel, mixing may be accomplished by end-over-end rocking in a sealed container. If preparing larger quantities of activated gel, use an overhead paddle stirrer. During the course of the reaction, the release of pyridyl-2-thione molecules will turn the solution a yellow color.

4. Wash the activated gel with at least 10 bed volumes of acetone:water (1:1) to remove excess reactants and reaction byproducts. Continue to wash thoroughly with water to remove the last traces of acetone. Finally, wash the support with 1-mM EDTA (at neutral pH) to protect against metal-catalyzed oxidation. Store the pyridyl disulfide-activated support at 4°C as a 50% slurry in water containing 1-mM EDTA and a preservative until use.

(B) Activation using NHS–PEGn–Pyridyl Disulfide Crosslinkers

PEG-containing heterobifunctional crosslinkers con-taining an amine-reactive NHS ester on one end and a pyridyl disulfide group on the other end may be used to form thiol-reactive groups by modification of amine-containing supports according to the following pro-tocol. Some optimization of the amount of crosslinker addition may have to be carried out to obtain the best density of pyridyldithiol groups on the support for a specific application. Refer to Chapter 18 for additional information on discrete PEG-based reagents. Other pyridyl disulfide-containing crosslinkers may also be used to activate an amine-support, such as SPDP or SMPT (Chapter 6, Sections 1.1 and 1.2).

1. Prepare 10 ml of a support containing an amine-terminal spacer using the methods described in the section on spacer arms in this chapter. The support should contain 10 to 50 μmoles –NH2 groups per ml of gel. Wash the support with water to remove storage solution components, if necessary, and drain to a wet cake.

2. In a fume hood, dissolve 0.56 g SPDP–dPEG4–NHS ester (also called PEG4–SPDP) (Quanta Biodesign or Thermo Fisher) in 2 ml of dry DMSO, DMF, or DMAC solvent. This amount will provide 100 μmoles of crosslinker per milliliter of gel during the reaction.

3. Add the moist gel cake from step 1 to 8 ml of reaction buffer (0.1-M sodium phosphate, pH 7.5) and mix to resuspend the beads.

4. With stirring, add the 2 ml of dissolved crosslinker prepared in step 2 to the stirring support slurry prepared in step 3.

5. React with constant mixing at room temperature for 1 h. Mixing may be done using an overhead paddle stirrer or by using end-over-end rocking in a sealed container.

6. Wash the activated gel thoroughly with water, 1-M NaCl, and water to remove excess reactants and reaction byproducts. Finally, wash the support with 1-mM EDTA (at neutral pH) to protect against metal-catalyzed oxidation of the disulfide groups. Store the pyridyl disulfide-activated support at 4°C as a



50% slurry in water containing 1-mM EDTA and a preservative until use.

(C) Activation using PDEAThe following protocol can be used to create reac-

tive pyridyl disulfide groups on an amine-reactive sup-port. The procedure uses the compound PDEA, which is 2-(2-pyridinyldithio)ethaneamine, containing a free pri-mary amine on one end and a pyridyldithiol group on the other end. PDEA may be obtained from GE Healthcare or synthesized by the reaction of 2 equivalents of 2,2′-dipyri-dyl disulfide (Aldrich) with 1 equivalent of cysteamine (or 2-mercaptoethylamine) in methanol according to the procedure of Li et al. (2008). The preparation of an amine-reactive support can be accomplished using any of the protocols previously described in this chapter; however, do not use an aldehyde-containing support to couple PDEA through reductive amination, because the presence of a reducing agent also will reduce the pyridyl disulfide groups and render the support inactive.

1. Prepare 10 ml of an amine-reactive support material according to the methods described in this chapter (e.g., CDI or an NHS ester-activated agarose). If the support is stabilized in an organic solvent (e.g., acetone), just before use filter off the excess solution using a sintered glass filter suspended in a vacuum filter flask within a fume hood. Draw a gentle vacuum until the excess solvent stops dripping from the filter while at the same time breaking up the support into finely divided pieces using a spatula, but do not allow it to dry. Release the vacuum.

2. In a fume hood, prepare a 20-mM PDEA solution in the appropriate solvent for coupling to the amine-reactive support of choice. For instance, if using a CDI- or NHS ester-activated support, to obtain maximal coupling yields dissolve 44 mg of PDEA in 10 ml of nonaqueous solvent, such as DMSO or DMF. Add to this solution, 2 mole equivalents (over the amount of PDEA added) of an organic base such as DIEA, TEA, or DMAP to catalyze the reaction by accepting the protons that are generated during the course of the reaction. Note: PDEA may also be coupled to amine-reactive supports in aqueous solution. The buffer composition and pH should be determined based on the suggested methods associated with the amine-reactive chemistries described earlier in this chapter. For aqueous coupling, mix the finely divided pieces of activated gel from step 1 with the PDEA buffer solution. The PDEA concentration may be increased for aqueous coupling to account for potential hydrolysis of reactive groups on the support. For instance, PDEA may be dissolved at 80-mM concentration in

50-mM sodium borate, pH 8.5, for amine-reactive immobilization methods compatible with this environment (i.e., for use with CDI- or NHS ester-activated supports).

3. With stirring and in the fume hood, mix the activated support material with the PDEA solution to resuspend it as a uniform slurry.

4. React with mixing for 1 h at room temperature using an overhead paddle stirrer or end-over-end rocking in a sealed container.

5. Wash the thiol-reactive support with solvent to remove excess reactants (at least 10–20 bed volumes). Wash the support into water by using sequentially increasing concentrations of water-in-solvent (e.g., 30% water, 70% water) until 100% water washes are used to completely remove the solvent. Note: If an aqueous reaction was done, then instead of washing with solvent wash with water, 1-M NaCl, and water to remove excess reactants. Finally, wash the support with 1-mM EDTA (at neutral pH) to protect against metal-catalyzed oxidation during storage. Store the pyridyl disulfide-activated support at 4°C as a 50% slurry in water containing 1-mM EDTA and a preservative until use.

LIGAND COUPLING TO PYRIDYL DISULFIDE-ACTIVATED SUPPORTS

The coupling of thiol-containing ligands to pyridyl-disulfide-activated supports can be performed over a broad pH range of approximately pH 4 to 8 with excel-lent results. The following immobilization protocols are generalized methods for coupling cysteine-containing proteins, peptides, or other thiol-containing molecules. The methods may also be used with proteins modified to contain –SH groups through the use of thiolation reagents (Chapter 2, Section 4.1). Some optimization may have to be carried out to determine a ligand den-sity that is best for a particular affinity chromatogra-phy application. Preparing a series of small batches using different concentrations of ligand in each reac-tion medium will produce a range of ligand densities on the support for evaluation. In addition, after ligand coupling do not attempt to block excess reactive groups with a thiol-containing compound such as cysteine, as this may also cleave off any immobilized ligands.

(A) Coupling of Reduced IgG or F(ab’)2 Fragments1. Dissolve an IgG antibody or F(ab’)2 fragment at

a concentration of 1 to 10 mg/ml in 1 ml of 0.1-M sodium phosphate, 0.15-M NaCl, 10-mM EDTA, pH 7.5 (coupling buffer). The coupling buffer should be purged with nitrogen and degassed under vacuum to remove oxygen, which may oxidize the thiols and prevent coupling.



2. Add a reducing agent, such as DTT, 2-mercapto-ethylamine (2-MEA), 2-mercaptoethanol (2-ME), or tris(carboxyethyl)phosphine (TCEP), to the antibody solution to give a final concentration of at least 5-mM. A highly concentrated stock solution of the reducing agent first may be prepared in coupling buffer and then a small aliquot of that solution added to the antibody to obtain the final desired concentration. Mix to dissolve and incubate for 1.5 h at 37°C. Note: Higher concentrations of the reducing agent may be used (typically ≤50-mM), but the higher the concentration, the more likely the reduction of disulfides between the heavy and light chains, which may disrupt the three-dimensional structure of the antibody and destroy antigen binding capability. Optimization of the reductant concentration may have to be done to ensure the best performance of the resultant immunoaffinity support. An alternative to the use of a reducing agent to generate free thiols is to use a thiolation reagent to first modify the amines on an antibody to contain thiols. Various procedures on the use of thiolation reagents with proteins and antibodies can be found elsewhere in this book (Chapter 2, Section 4.1).

3. Remove excess reducing agent from the antibody solution by desalting using size exclusion chromatography or by dialysis. During the chromatography or dialysis operation, use coupling buffer to equilibrate and elute the protein through the gel filtration support or as the dialyzing solution. To desalt the 1-ml sample size, a gel filtration column should consist of at least 10 ml of a support having a molecular weight exclusion limit of no more than 5 to 10 kDa to ensure that the protein will come through first in the void volume and be separated from the salt peak containing the reducing agent. A similarly sized dialysis membrane having a size exclusion limit of 5 to 10 kDa also is appropriate. Dialyze the antibody with numerous changes of the outside dialysis solution to ensure that the concentration of the reducing agent is decreased down to levels far below that of the reduced antibody. Complete removal of the reducing agent is essential to eliminate competition when coupling the reduced antibody to the pyridyl disulfide-activated support. Pool the fractions containing desalted protein from the gel filtration column or recover the dialyzed protein from the dialysis device. The solution may be concentrated if necessary using a centrifugal concentrator to approximately 1 ml to maintain the desired concentration level of antibody in the coupling reaction.

4. Wash 1 ml of a pyridyl disulfide-activated support with several bed volumes of water to remove storage solution components and then with two bed volumes

of coupling buffer. Washing may be performed using spin columns or by using a small drip column. Drain to a moist cake.

5. Mix the washed pyridyl disulfide-activated gel with the reduced antibody solution and stir at room temperature (or 4°C) overnight or for at least 18 h. Mixing may be achieved by end-over-end rocking in a sealed container. If the antibody is sensitive to extensive stirring or rocking, the mixture may be mixed for 2 h and then left to sit with periodic resuspension and mixing every hour or so to maintain a somewhat homogeneous mixture of ligand throughout the resin.

6. Wash the support with several bed volumes of coupling buffer to remove most of the not-coupled antibody. The washes may be analyzed versus the initial concentration of the antibody solution before coupling to determine the amount immobilized.


(B) Coupling Thiol-Containing Small Ligands1. Wash 10 ml of a pyridyl disulfide-activated support

with water and then place into coupling buffer using a sintered glass filter funnel suspended in a suction filter flask (coupling buffer: 0.1-M sodium phosphate, 0.15-M NaCl, 10-mM EDTA, pH 7.5). The coupling buffer should be purged with nitrogen and degassed under vacuum to remove oxygen, which may oxidize the thiols and prevent coupling.

2. Dissolve a thiol-containing ligand (such as a cysteine-containing peptide) in 10 ml of coupling buffer at a concentration of at least 2 to 5 mg/ml. Determination of the optimal level of ligand concentration may be done experimentally by running a series of small reactions at different concentrations and determining which preparation performs best in the intended affinity chromatography application.

3. Add the ligand solution to the washed pyridyl disulfide-activated resin and mix to thoroughly resuspend the support in the solution. React overnight or for at least 18 h at room temperature (or 4°C) with constant mixing.

4. Wash the gel with several bed volumes of coupling buffer to remove most of the excess uncoupled ligand. The amount of ligand not coupled may be determined by measuring the volume and concentration of the ligand in the pooled washes. This may be done using a protein assay or spectrophotometrically if the ligand has a characteristic spectral signature. The amount



of ligand that coupled to the matrix then may be determined by difference. Drain the gel to a wet cake.


TNB–Thiol ActivationEllman’s reagent (5,5′-dithiobis(2-nitrobenzoic acid),

or DTNB, long has been used as a chromogenic agent for the determination of free thiol groups, especially cysteine residues in proteins (Chapter 2, Section 4.1) (Ellman, 1958; Riddles et al., 1979, 1983; Watabe et al., 1982). The compound reacts with thiol groups in a disulfide exchange process to generate the chromogenic leaving group 5-thio-2-nitrobenzoic acid (TNB). The yellow color generated by the TNB anion at 412 nm can be used to quantify the amount of thiol groups present (ε = 14,150 at pH 8.0).

Ellman’s reagent also can be used to create thiol-reactive groups on affinity supports for the coupling of sulfhydryl-containing ligands (Jayabaskaran et al., 1987). Much like the reaction of 2,2′-dipyridyl disulfide discussed in the previous section, DTNB can be used to activate thiol-containing chromatography supports to contain reactive TNB–thiol groups (Figure 15.78). The TNB–thiol groups then can be used to immobilize thiol-containing proteins or other ligands through a disulfide exchange process leading to the formation of a disulfide linkage with the affinity ligand (Figure 15.79). The use of Ellman’s reagent in this process provides advantages over the pyridyl disulfide chemistry in that the reactions of the TNB group can be followed visu-ally and spectrophotometrically. The activation of a

thiol-containing support proceeds with loss of one chro-mogenic TNB group per thiol, so the reaction can be quantified by measuring the amount of color released during the process. In addition, the immobilization of a thiol-containing protein or molecule can be quantified through the number of TNB groups generated during the coupling reaction.

The requirement for production of a TNB-activated support is to first prepare a spacer arm derivative that terminates in a thiol group. This can be done using the techniques discussed in this chapter on the preparation of spacer arm derivatives. Perhaps one of the easiest routes to accomplish this process is to use an amine-containing support (such as DADPA–agarose or MANEA–agarose) and modify it with Traut’s reagent to spontaneously form a small spacer arm, which opens up to a free thiol on its terminus (Chapter 2, Section 4.1). This intermediate can then be immediately reacted with DTNB to create the activated TNB–thiol support (Figure 15.80).

PREPARATION OF A TNB–THIOL-ACTIVATED SUPPORT

The following protocol requires the prior preparation of an amine-containing support for the initial modifica-tion with a thiolation reagent to create thiols (see sec-tion on spacer arms in this chapter).

1. Wash 100 ml of an amine-containing support (such as DADPA–agarose or MANEA–agarose) with water to remove any storage components (at least 3 bed volumes) and then with 0.1-M sodium phosphate, 0.15-M NaCl, 10-mM EDTA, pH 8.0 (modification buffer). Use a sintered filter funnel suspended in a suction filter flask to facilitate the washing steps, while applying a gentle vacuum between each wash solution addition to accelerate flow through the resin. Drain to a wet cake.

Thiol-containingsupport

+

TNB-activated support

SH

S

SS

O

HO

N+O

- OO

OH

N+

O

O-

DTNB;5,5'-Dithiobis(2-nitrobenzoic acid)

SO

OH

N+O

O-

HSO

OH

N+O

O-5-Thio-2-nitrobenzoicacid (TNB); A 412 nm

FIGURE 15.78 Activation of a thiol-containing matrix with DTNB.



2. Prepare 100 ml of a 20-mM 2-iminothiolane (Traut’s reagent) solution in modification buffer and add the gel cake to it with stirring.

3. React for 1 h at room temperature with constant stirring using an overhead paddle stirrer or end-over-end rocking in a sealed container.

4. Quickly wash the support with several bed volumes of modification buffer to remove most

of the reactant and then drain to a wet cake. Note: Do not store the thiolated support at this stage, because 2-iminothiolane modified species may undergo a recyclization reaction to an inactive state with the thiol group unavailable for further modification (refer to other sections on Traut’s reagent within this book for additional information).

Ligand coupledthrough thioether bond

S

HSO

OH

N+O

O-5-Thio-2-nitrobenzoicacid (TNB); A 412 nm

TNB-activated support

SS

O

OH

N+O

O-


+S

HS

FIGURE 15.79 Coupling of a thiol-containing ligand to a TNB-thiol-activated support.

O

HN

NH2


primary amines

+ O

HN

Thiolated supportcontaining free –SH group

O

HN

Traut's reagent

TNB-thiol support

SS

O

HO

N+O

- OO

OH

N+

O

O-

DTNB

+

S NH2Cl

SHNH2Cl

NH

NH2Cl

NH

SS

O

OH

N+O

O-

FIGURE 15.80 Reaction of Traut’s reagent with MANEA–agarose and activation with DTNB.



5. Prepare 100 ml of a 20-mM DTNB (Ellman’s reagent) solution in modification buffer and add the gel cake to it with stirring.

6. React for 1 h at room temperature with constant mixing. During the reaction, the chromophoric TNB anion will be released and generates a yellow color within the gel slurry. This is indicative of a successful activation process.

7. Drain the activated gel of excess reaction solution and wash with an additional 2 bed volumes of modification buffer. These initial washes may be collected and pooled to determine the activation level of the support by measuring the absorbance of the released TNB anion at 412 nm (ε = 14,150 at pH 8.0). Continue to wash the activated support thoroughly with water, 1-M NaCl, and water to remove excess reactants and reaction byproducts. Finally, wash the support with 1-mM EDTA (at neutral pH) to protect against metal-catalyzed oxidation of the disulfide groups. Store the TNB–thiol-activated support at 4°C as a 50% slurry in water containing 1-mM EDTA and a preservative until use.

LIGAND COUPLING TO TNB–THIOL-ACTIVATED SUPPORTS

The following protocols describe the immobiliza-tion of a thiol-containing reduced antibody or a thiol-containing small ligand (such as a peptide containing a cysteine group) to a TNB–thiol-activated resin. The methods may also be used with proteins that have been modified to contain –SH groups through the use of thiolation reagents (Chapter 2, Section 4.1). Some optimization may have to be done to determine a ligand density that is best for a particular affinity chro-matography application. Preparing a series of small batches using different concentrations of ligand in each reaction medium will produce a range of ligand den-sities on the support for evaluation. In addition, after ligand coupling do not attempt to block excess reac-tive groups with a thiol-containing compound such as cysteine, as this may also cleave off any immobilized ligands.

(A) Coupling of Reduced IgG or F(ab’)2 Fragments1. Dissolve an IgG antibody or F(ab’)2 fragment at

a concentration of 1 to 10 mg/ml in 1 ml of 0.1-M sodium phosphate, 0.15-M NaCl, 10-mM EDTA, pH 8.0 (coupling buffer). The coupling buffer should be purged with nitrogen and degassed under vacuum to remove oxygen, which may oxidize the thiols and prevent coupling.

2. Add a reducing agent, such as DTT, 2-mercapto-ethylamine (2-MEA), 2-mercaptoethanol (2-ME), or tris(carboxyethyl)phosphine (TCEP), to the antibody

solution to give a final concentration of at least 5-mM. A highly concentrated stock solution of the reducing agent may first be prepared in coupling buffer and then a small aliquot of that solution added to the antibody to obtain the final desired concentration. Mix to dissolve and incubate for 1.5 h at 37°C. Note: Higher concentrations of the reducing agent may be used (typically ≤50-mM), but the higher the concentration the more likely will be the reduction of disulfides between the heavy and light chains, which may disrupt the three-dimensional structure of the antibody and destroy antigen binding capability. Optimization of the reductant concentration may have to be done to ensure the best performance of the resultant immunoaffinity support. An alternative to the use of a reducing agent to generate free thiols is to use a thiolation reagent to modify the amines on an antibody to contain thiols. Various procedures on the use of thiolation reagents with proteins and antibodies can be found elsewhere in this book (Chapter 20, Section 1.1).

3. Remove excess reducing agent from the antibody solution by desalting using size exclusion chromatography or by dialysis. During the chromatography or dialysis operation, use coupling buffer to equilibrate and elute the protein through the gel filtration support or as the dialyzing solution. To desalt the 1-ml sample size, a gel filtration column should consist of at least 10 ml of a support having a molecular weight exclusion limit of no more than 5 to 10 kDa to ensure that the protein will come through first in the void volume and be separated from the salt peak containing the reducing agent. A similarly sized dialysis membrane having a size exclusion limit of 5 to 10 kDa is also appropriate. Dialyze the antibody with numerous changes of the outside dialysis solution to ensure a decrease in the concentration of the reducing agent down to levels far below that of the reduced antibody. Complete removal of the reducing agent is essential to eliminate competition when coupling the reduced antibody to the TNB–thiol-activated support. Pool the fractions containing desalted protein from the gel filtration column or recover the dialyzed protein from the dialysis device. The solution may be concentrated if necessary using a centrifugal concentrator to approximately 1 ml to maintain the desired concentration level of antibody in the coupling reaction.

4. Wash 1 ml of a TNB–thiol-activated support with several bed volumes of water to remove storage solution components and then wash with two bed volumes of coupling buffer. Washing may be carried out using spin columns or by using a small drip column. Drain to a moist cake.



5. Mix the washed TNB–thiol-activated gel with the reduced antibody solution and stir at room temperature (or 4°C) for 2 to 4 h. Mixing may be carried out by end-over-end rocking in a sealed container (which can consist of the sealed column used for washing). If the antibody is sensitive to extensive stirring or rocking, the mixture may be mixed for 2 h and then left to sit with only periodic resuspension and mixing every 30 min or so to maintain a somewhat homogeneous mixture of ligand throughout the resin.

6. Wash the support with several bed volumes of coupling buffer to remove most of the not-coupled antibody. The washes may be analyzed versus the initial concentration of the antibody solution before coupling to determine the amount immobilized by difference. During the course of the reaction, TNB groups will also be released as the thiol-containing antibody has coupled; therefore, measurement of the absorbance of this anion at 412 nm can provide an accurate determination of the amount of ligand coupled.


(B) Coupling Thiol-Containing Small Ligands1. Wash 10 ml of a TNB–thiol-activated support with

water and then add into coupling buffer using a sintered glass filter funnel suspended in a suction filter flask (coupling buffer: 0.1-M sodium phosphate, 0.15-M NaCl, 10-mM EDTA, pH 8.0). The coupling buffer should be purged with nitrogen and degassed under vacuum to remove oxygen, which may oxidize the thiols and prevent coupling.

2. Dissolve a thiol-containing ligand (such as a cysteine-containing peptide) in 10 ml of coupling buffer at a concentration of at least 2 to 5 mg/ml. Determination of the optimal level of ligand concentration may be achieved experimentally by running a series of small reactions at different concentrations and determining which preparation performs best in the intended affinity chromatography application.

3. Add the ligand solution to the washed TNB–thiol-activated resin and mix to thoroughly suspend the support in the solution. React for 2 to 4 h at room temperature (or 4°C) with constant mixing.

4. Wash the gel with several bed volumes of coupling buffer to remove most of the excess uncoupled ligand. The amount of ligand not coupled may be determined by measuring the volume and concentration of the ligand in the pooled washes. This may be done using a protein

assay or spectrophotometrically if the ligand has a characteristic spectral signature. The amount of ligand that coupled to the matrix may then be determined by the difference. Alternatively, monitoring the amount of the TNB anion released during the coupling reaction through its absorbance at 412 n m can provide an indirect, but accurate measurement of the degree of ligand coupling. Drain the gel to a wet cake.


2.3. Hydroxyl-Reactive Immobilization Methods

This section describes the immobilization of hydroxyl-containing ligands to chromatography sup-ports. Activated supports that are able to couple effec-tively with hydroxyl-containing ligands must be able to react with the hydroxyl groups while avoid-ing substantial inactivation by hydrolysis in aqueous solution. The following activation methods perform particularly well in this regard for coupling sugars, car-bohydrates, polysaccharides, glycans, and other mol-ecules containing one or more hydroxyl groups. One should keep in mind that in addition to these methods meant to first activate the matrix and then attach the ligand, hydroxyl-containing ligands could also be ini-tially activated in solution with any of the activating reagents described in this chapter suitable for hydrox-yls and then in a reverse manner reacted with a sup-port material that contained the appropriate functional group for coupling to the activated ligand intermediate. In other words, a hydroxyl-containing molecule might be activated in organic solution with a limiting mole quantity of DSC, for example (see previous section on DSC activation, this chapter), and then the resultant NHS carbonate reactive group formed on the ligand could be coupled to an amine-containing support mate-rial to form a carbamate linkage. This strategy of revers-ing the normal sequence of support activation followed by ligand coupling to be instead ligand activation fol-lowed by coupling to a support can be performed with many organic-soluble small molecules containing a hydroxyl group. This alternative strategy therefore may open up many more options for designing a reaction sequence leading to the desired immobilized ligand.

Epoxide ActivationAn epoxide group is a small three-member heterocy-

clic ring containing two carbon atoms and one oxygen. The combination of ring strain and the electron with-drawing effects of the oxygen atom cause the carbon



atoms to carry a partial positive charge and therefore be good sites for potential nucleophilic attack. An epoxide reacts with a nucleophile through a ring opening pro-cess, which yields a bond to the nucleophilic group on one carbon and the formation of a hydroxyl group on the adjacent carbon as the epoxy ring opens. For epox-ides that exist on terminal carbon atoms, the reaction with a nucleophile causes covalent linkage with the pri-mary carbon atom and hydroxyl formation on the adja-cent secondary carbon atom.

Depending on the pH and conditions of the reac-tion medium, epoxides can be made to react effec-tively with thiols, amines, or hydroxyl (even phenolic) groups (Figure 15.81). Similar to the relative reactiv-ity of vinyl sulfone-activated supports described pre-viously, epoxides efficiently react with thiols in the pH range of 7.5 to 8.5, amines react within the range

of pH 9 to 11 (Ghazi et al., 2005), and hydroxyl groups react at highly alkaline pH values (pH > 11). The reac-tions may be accelerated with heating (i.e., ≥37°C), which often is done during the activation of a hydroxyl-containing support with a bis-epoxide compound and during subsequent coupling reactions with carbohy-drate hydroxyl groups. Epoxy groups also are known to react under certain conditions with carboxylic acids to form ester linkages. This reaction is important in many polymer resins that are designed to crosslink and form coatings, adhesives, moldings, and other synthetic sol-ids for industrial and commercial applications (Blank et al., 2001). Epoxy ester formation has also been inves-tigated with amino acids as a mechanism in the prepa-ration of crosslinked collagen-based materials, and it has found to occur best at slightly acidic pH (Zeeman, 1998). However, the reaction rate of carboxylate groups with epoxides is much slower than that of the reactions with thiols, amines, or hydroxyl groups under the rec-ommended pH conditions described in this section. It is likely that if ester formation occurs with biological ligands that it is a minor product compared to the reac-tions commonly used with epoxy-activated supports.

Homobifunctional epoxide crosslinker compounds (bisoxiranes) are often used to activate hydroxyl-containing chromatography supports to result in the production of reactive epoxy groups for the immobili-zation of hydroxylic ligands. Perhaps the reagent most commonly used for this purpose is 1,4-butanediol diglycidyl ether (Sundberg and Porath, 1974), which contains a relatively hydrophilic 12-atom spacer group with epoxides at each end. At alkaline pH and in large excess, one end of this compound can be made to react with the hydroxyls on a matrix to create an ether link-age while forming a hydrophilic spacer which ter-minates in the second epoxide group for subsequent ligand immobilization (Figure 15.82).

The coupling of protein affinity ligands onto epox-ide-activated supports can be significantly enhanced by the addition of certain salts to the reaction medium (Murthy and Moudgal, 1986; Wheatley and Schmidt, Jr., 1993, 1999). In particular, salts that increase the interactions of hydrophobic molecules in aqueous solu-tion appear to promote protein adsorption onto the activated support prior to covalently reacting with the epoxy groups. Lyotropic salts such as sodium or potas-sium sulfate, sodium or potassium phosphate, and even ammonium sulfate can be used to drive soluble proteins toward the surface of the particles by a salt-ing out effect, thus forcing them into proximity with the reactive groups. Surprisingly, even the presence of ammonium ions in ammonium sulfate salts does not interfere with the reaction with amines on proteins. Using ammonium sulfate concentrations of 0.4 to 2.5-M, most proteins will couple to the epoxide support in




Thioether linkage


Ether linkage

Incr

easi

ng p

H

Epoxide-activated support

O

HO

O

OOH

O

Ester linkage

SOH

HN

OH

OHO

HS

HO

H2N

Carboxyl-containingligand

FIGURE 15.81 Reaction of an epoxide support with a thiol-, amine-, hydroxyl-, or carboxylate-containing compound.



yields of 95 to 100% in 20 h at room temperature. The other activated support that has this tendency to be enhanced for coupling proteins when in the presence of a lyotropic agent is the azlactone activation chemis-try, described previously in this chapter. Presumably, the lyotropic-induced deposition of proteins onto the epoxide-activated surface occurs first, followed by the nucleophilic attack of protein amines onto the carbon of the epoxide ring, which causes covalent coupling.

Even the immobilization of some small affin-ity ligands onto epoxide-activated supports can be enhanced by the addition of a lyotropic salt to the cou-pling reaction. Bauer-Arnaz et al. (1998) found that vari-ous S-alkyl glutathione derivatives could be coupled in approximately twice the yield if a high concentra-tion of potassium phosphate (2-M) was present in the reaction buffer. In this case, the coupling reaction was performed at pH 10.5, which indicates that even amine-containing small ligand coupling can be enhanced with salt if the compound structure contains hydrophobic regions that can interact with the activated support prior to covalent binding.

Mateo et al. (2002) developed salt-induced immo-bilization methods using the epoxide-activated sup-port Sepabeads-EP. The coupling and stabilization of the enzyme penicillin G acylase (PGA) from E. coli was accomplished using an initially high concentra-tion of sodium phosphate buffer (1-M) at pH 7. The effect of the salt to push the enzyme molecules toward the matrix surface near the epoxide active groups resulted in efficient immobilization even at neutral pH. Although the reaction took 24 h to go to completion, the result was an increase of around 10,000-fold in stabiliza-tion of the enzyme compared to the soluble form.

Grazú et al. (2003) used an interesting combination of thiol/disulfide exchange coupling and epoxide reac-tions for enzyme immobilization to enhance the yield of epoxy coupling without the use of a lyotropic salt (see previous sections on pyridyl disulfide and TNB–thiol activation, this chapter). On a highly activated

epoxide support, they reacted a limited amount of DTT with it to obtain a partially thiolated matrix, which contained both thiol and epoxide groups. Next, an enzyme was thiolated with SPDP followed by reduc-tion and the thiols on the support were activated using 2,2′-dipyridyl disulfide to provide reactive pyridyldi-thiol groups. Mixture of the thiolated enzyme with the activated support resulted in the rapid formation of disulfide linkages with the thiol groups created on the protein followed by continued reaction with the epox-ide groups to give a multi-point attachment with each enzyme.

Epoxy activation and coupling has been extensively used to couple metal chelating compounds to particles of all types for immobilized metal affinity chromatogra-phy (IMAC) applications (Novotna et al., 2010). Often, an organic chelating molecule contains a secondary amine, such as in the example of iminodiacetic acid, which can be effectively coupled to a chromatography support by reaction with an epoxide (Figure 15.83). In addition, the coupling of glutathione is often done using epoxy-activated supports, because the most effec-tive site of immobilization on the molecule is through its thiol group, which allows the ligand to interact with glutathione-S-transferase (GST) fusion proteins to cap-ture and purify them after recombinant expression (Simons and Vander Jagt, 1977; Smith and Johnson, 1988; Stahl et al., 2003).

Due to the versatility of epoxy-activated supports in coupling thiol-, amine-, or hydroxyl-containing ligands, the method has become a popular choice for immobi-lizing a wide range of affinity molecules. In addition, the activated support is extremely stable to hydroly-sis even when stored in aqueous solution (neutral pH) for extended periods. The activation of chromatogra-phy supports, nonporous particles, and surfaces with the compound 1,4-butanediol diglycidyl ether creates a long spacer arm linking any potential ligand, which is also hydrophilic enough to minimize nonspecific binding.

OO

OO

1,4-Butanediol diglycidyl ether

OH


+

OOH

OO

O

Epoxy-activated support

OH-

FIGURE 15.82 Activation of hydroxylic supports with 1,4-butanediol diglycidyl ether.



PREPARATION OF AN EPOXY-ACTIVATED SUPPORT

The following protocol describes the activation of a hydroxyl-containing support with the bis-epoxide com-pound 1,4-butanediol diglycidyl ether under highly alkaline conditions. All operations should be done in a fume hood while wearing the appropriate personal pro-tective equipment. Avoid contact with all solutions and avoid breathing vapors of the epoxide compound or the solvents used during the operations.

1. Wash 100 ml of a hydroxyl-containing support material (i.e., Sepharose (agarose), Toyopearl, or Trisacryl) with several bed volumes of water to remove storage solution and preservatives. The washing may be done using a sintered glass filter funnel suspended in a suction filter flask. Pull a gentle vacuum on the flask to facilitate the removal of the solutions from the gel after each addition of wash. Drain to a moist cake. Note: When activating agarose supports with a bis-epoxy compound, it is common to start with a non-crosslinked support (e.g., 4% agarose), because reactions with both ends of the epoxide will occur to some extent and form a crosslinked matrix.

2. Suspend the gel in 75 ml of 0.6-N NaOH (caustic!) containing 150 mg of sodium borohydride dissolved

into it. Transfer the gel suspension into a three-necked, round-bottom flask and stir using an overhead paddle stirrer in the fume hood. Place the round-bottom flask in an open container into which water can be added to control the temperature of the reaction. Put a thermometer into one of the three necks of the flask to monitor the temperature throughout the activation reaction. Add water at room temperature to the surrounding bath up to the level of the inner gel slurry and have ice on hand if necessary to control any exothermic tendencies.

3. Slowly add to the stirring gel suspension 75 ml of 1,4-butanediol diglycidyl ether over a period of about 20 min. Monitor the temperature and add ice to the surrounding water bath if required to maintain a constant ambient temperature of about 20 to 25°C. There may be an exothermic temperature spike during the early stages of the reaction; however, do not cool the reaction below the recommended temperature range, as it will slow the activation process.

4. Stir the reaction for 8 to 10 h or overnight at room temperature.

5. Transfer the gel slurry to the filter funnel and wash with 4 bed volumes of water to remove the NaOH solution and the borohydride. The excess epoxide


O

+

OHN

OO

HS

NH

NH2

OO

O

Glutathione

Immobilized glutathionethrough thioether bond

OHOHN

O

O

SNH

NH2

OO

O


O

+OH

HN

O

O

O

O

Iminodiacetic acid

O

O

OO

N

Immobilized iminodiaceticacid through tertiary amine bond

FIGURE 15.83 Immobilization of iminodiacetic acid and glutathione on epoxy-activated supports.



compound is more difficult to remove using only water washes. For this reason, sequentially wash the gel with increasing concentrations of acetone (or ethanol) (e.g., 30%, 70%) until 100% solvent is used. Continue to wash with 100% solvent for at least 10 bed volumes to remove the remaining traces of excess epoxide. Next, sequentially wash back into water by reversing the concentrations of solvent used previously. Finally, wash with 10 bed volumes of water to remove the last traces of solvent. Store the epoxy-activated support at 4°C as a 50% slurry in water and containing a preservative until use.

LIGAND COUPLING TO EPOXY-ACTIVATED SUPPORTS

The following protocols represent some common methods of coupling ligands to epoxy-activated sup-ports. In the first example, a disaccharide (lactose) is immobilized through its hydroxyl groups onto epoxy-activated agarose at high pH. Other sugars or carbohy-drates, including polysaccharides and glycans, may be coupled using similar procedures. In the second exam-ple, a protein is immobilized onto an epoxy-activated support at near physiological pH using a buffer con-taining a lyotropic agent to push the protein molecules into close proximity with the reactive groups, and thus facilitate efficient coupling while avoiding higher pH conditions to drive the reaction.

(A) Coupling of Carbohydrates1. Wash 100 ml of an epoxy-activated support with

several bed volumes of water to remove any storage solution and preservatives. Use a sintered glass filter funnel suspended in a vacuum filtration flask to facilitate the washing steps. Drain the gel to a moist cake.

2. Prepare a lactose ligand solution by dissolving 15 g of lactose in 100 ml of 0.1-N NaOH (makes a 0.438-M solution). Use care in handling the caustic NaOH solution and use the appropriate personal protective equipment during all operations.

3. Add the washed gel cake to the ligand solution with stirring and transfer the reaction slurry to a three-necked, round-bottom flask. Put the flask in a heating mantle and use an overhead paddle stirrer to mix the resin during the reaction. Add a thermometer to one of the openings in the flask and heat the reaction mixture with constant stirring to 40°C.

4. Stir the reaction for 24 h at 40°C.5. Wash the support with several bed volumes of

water to remove most of the reaction medium and then wash with 0.1-M sodium bicarbonate, pH 8.0, followed by an extensive water wash to remove the last traces of buffer and reactants. The immobilized lactose support can be stored until use at 4°C as a 50% slurry in water containing a preservative.

(b) Coupling of ProteinsProteins containing cysteine thiol groups or amines

may be immobilized onto epoxy-activated supports in the pH range of 7.0 to 11, with pH 7.5 to 8.5 being most effective for –SH groups and pH 9 to 11 optimal for lysine or N-terminal amines (Ghazi et al., 2005). However, if a lyotropic salt is added to the reaction medium, then protein coupling to amines can be made to occur with excellent yield at pH 7.0 to 8.0, which is much more amenable to maintain protein stability. The following protocol describes the immobilization of a protein at mildly alkaline pH using the lyotropic salt sodium sulfate. Some optimization of the level of salt concentration in the reaction medium may have to be carried out for certain proteins to prevent precipitation while still promoting the best possible matrix-protein interactions leading to efficient reaction rates.

1. Wash 10 ml of an epoxy-activated support with several bed volumes of water to remove any storage solution and preservatives. Use a small sintered glass filter funnel suspended in a vacuum filtration flask to facilitate the washing steps. Drain the gel to a moist cake.

2. Prepare 10 ml of a protein ligand solution at a concentration of 1–20 mg/ml in coupling buffer. The coupling buffer may be formulated as 0.1-M sodium phosphate, pH 7.5, containing 0.5-M sodium sulfate as the lyotropic agent. Other lyotropic salts may also be used. For example, an alternative coupling buffer composition that was used with success by Mateo et al. (2002) to immobilize enzymes consisted of 1-M sodium phosphate, pH 7.0, in which the sodium phosphate functioned both as a buffer for pH stabilization and as the lyotropic agent. Despite the effect of a lyotropic agent to drive protein molecules toward the surface for reaction, some proteins may still require a higher-pH environment to efficiently couple. An alternative higher pH buffer is 0.1-M sodium carbonate, pH 9.0, containing 0.5-M sodium sulfate.

3. Add the washed epoxy-activated resin to the ligand solution with stirring.

4. React with constant mixing for at least 24 h at room temperature. Mixing may be done using an overhead paddle stirrer or by end-over-end rocking in a sealed container. Some proteins may require longer reaction times to reach maximal coupling yields for attachment to the support.

5. Wash the support with several bed volumes of coupling buffer and collect the washes. The amount of protein coupled may be determined by the difference between the initial amount of protein added to the reaction mixture and the total amount left in the not-coupled washings. Drain the support to a wet cake.

6. Block unreacted epoxide groups on the support using a small molecule containing a nucleophilic



group that will readily react with the leftover active sites. For instance, cysteine, ethanolamine, or glycine may be used for this purpose. Cysteine blocking will proceed faster than amine-containing blockers and at lower pH values, because of the increased reaction rate of epoxides with the highly nucleophilic thiol groups. However, avoid using thiol-containing compounds that are also strong disulfide reducing agents, such as DTT or 2-mercaptoethanol, if the protein that was immobilized is sensitive to reduction. Cysteine is much milder in this regard and can be used as a blocker without detrimentally affecting most proteins. Prepare at least a 0.1-M solution of the blocking agent in coupling buffer and mix with the washed resin.

7. React 2 to 4 h with mixing (for a thiol-containing blocking agent, such as cysteine) or for 24 h (if using an amine-containing blocking agent).

8. Wash the support with 10 bed volumes of coupling buffer without the lyotropic agent present (e.g., 0.1-M sodium phosphate, pH 7.5). Continue to wash with water, 1-M NaCl, and again with water to remove the last traces of all reactants and coupling buffer components. The immobilized protein can be stored until use at 4°C as a 50% slurry in water containing a preservative.

Divinyl Sulfone ActivationDivinyl sulfone (DVS) is a small bifunctional com-

pound that can be used to activate hydroxyl-containing supports for the immobilization of hydroxyl-containing ligands, such as sugars, carbohydrates, polysaccha-rides, glycans, and small organic molecules containing an available –OH group. The chemistry of DVS activa-tion and coupling is discussed in great detail in a pre-vious section in this chapter, including a protocol for the coupling of hydroxylic ligands; see Thiol-Reactive Immobilization Methods for an overview and protocols for ligand immobilization.

Trichloro-s-Triazine ActivationThe activation of hydroxyl-containing supports with

trichloro-s-triazine (TsT) was presented in detail in the section on Amine-Reactive Immobilization Methods discussed earlier in this chapter. The cyanuric chloride reactive groups on a TsT-activated support can be used to immobilize thiol-, amine-, or hydroxyl-containing ligands with high efficiency. The coupling of hydroxyl-containing sugars, carbohydrates, polysaccharides, gly-cans, or other organic ligands containing hydroxyls can be done using a high pH environment (pH > 11), essen-tially using the same conditions that are recommended for the coupling of triazinyl dyes to hydroxyl matrices (Hermanson et al., 1992; Chamani et al., 2011).

LIGAND COUPLING TO TsT-ACTIVATED SUPPORTS

The following protocol assumes that 100 ml of a TsT-activated support has been prepared as described previ-ously in the section on Amine-Reactive Immobilization Methods.

1. Prepare 100 ml of a hydroxyl-containing ligand solution in 0.5-M sodium carbonate (no pH adjustment necessary) by dissolving the ligand at a concentration range of 0.1 to 0.5-M (for a small sugar molecule or disaccharide) or about 5 to 10 mg/ml (for larger polysaccharide or glycan ligands). Notes: The optimal concentration of a carbohydrate in the immobilization reaction may have to be determined by experimentation to obtain the best ligand loading for acceptable performance in the intended affinity chromatography application. For example, immobilized sugars used in the purification of lectins often are coupled at relatively low ligand levels (below the maximal coupling capacity of the gel), because a lower ligand density avoids the potential for multi-site binding with a lectin, which may result in high avidity interactions followed by difficulty in eluting the bound proteins from the support.

2. Wash 100 ml of a TsT-activated support into water and coupling buffer by suctioning off excess acetone storage solution and resuspending the gel into water. This can be done by pulling a gentle vacuum on the filter flask to filter off solvent while breaking up the support into small, finely divided pieces using a spatula, at the same time as being careful not to allow the matrix to dry out. Once the support is broken into small pieces, remove the vacuum and resuspend the gel in water with mixing. Continue to wash the support with water for at least 5 to 10 bed volumes to remove the solvent. Finally, wash the support with 2 to 3 bed volumes of coupling buffer (0.5-M sodium carbonate, no pH adjustment).

3. Add the washed gel cake to the ligand solution with stirring and transfer the reaction slurry to a three-necked, round-bottom flask. Put the flask in a heating mantle and use an overhead paddle stirrer to mix the resin during the reaction. Add a thermometer to one of the openings in the flask and heat the reaction mixture with stirring to 40°C. After 2 h of mixing, increase the temperature to 60°C with continued stirring.

4. Mix the reaction for 24 h at 60°C.5. Wash the support with several bed volumes of

water to remove most of the reaction medium and then wash with 0.1-M sodium bicarbonate, pH 8.0, followed by an extensive water wash to remove the last traces of buffer and reactants. The immobilized carbohydrate support can be stored until use at 4°C as a 50% slurry in water containing a preservative.



2.4. Carbonyl-Reactive Immobilization Methods

Chromatography supports can be modified with reactive groups to specifically couple to carbonyls on affinity ligands or proteins and form stable covalent bonds. These carbonyl groups may consist of alde-hydes, ketones, or sometimes even carboxylates on ligands that can be targeted under the right conditions to covalently link with certain reactive groups present on support materials. The following sections describe the principal methods of activating a support to couple with carbonyl-containing ligands. The reactive interme-diates prepared in this section are somewhat unique, however, in that they typically do not spontane-ously form covalent bonds with the functional groups encountered on most biomolecules. The hydrazide, aminooxy, and amine derivatives of supports described in this section will not immediately react with large complex macromolecules like proteins, nucleic acids, lipids, and most polysaccharides found within cells or living organisms. In this regard, they have more selec-tive reactivity than most of the other electrophilic active groups described in this chapter for the immobiliza-tion of biomolecules. However, given the right circum-stances where a ligand contains an aldehyde, ketone, or in some instances a carboxylate, the creation of these active intermediates can facilitate site-directed coupling to discrete locations on molecules that some of the other methods cannot accomplish.

Hydrazide or Hydrazine Supports for Coupling Aldehydes, Ketones, or Carboxylates

A hydrazine or hydrazide group is able to react with an aldehyde or ketone to form a dehydration prod-uct, called a hydrazone, which is a type of Schiff base having a double bond between the carbon atom of the original carbonyl group and the terminal hydrazino nitrogen. This product represents a more stable Schiff base than that formed between an amine and a car-bonyl group (standard imine), because a hydrazone is less susceptible to hydrolysis back to the starting mate-rials. In many cases, the hydrazone bond created with a carbonyl-containing ligand is strong enough not to need reduction with sodium cyanoborohydride to sta-bilize the linkage, as opposed to amino Schiff bases which almost always require reduction to prevent leakage. The hydrazone electrons can delocalize due to the electronegativity of the hydrazide carbonyl oxy-gen and therefore it stabilizes the bond much better than the linkage between an amine and an aldehyde group. Particularly, if there is a potential for more than one hydrazone attachment point with a ligand, such as in the immobilization of glycoconjugates, the likeli-hood of ligand leakage becomes negligible. In certain cases, however, hydrazone bonds should be reduced

to prevent the possibility of slow ligand leaching over time. This is especially true if the point of attachment between the support and the ligand consists of only a single hydrazone bond.

The use of hydrazides or hydrazines created on a chromatography matrix to immobilize carbonyl-containing ligands was developed specifically to couple glycoproteins through their carbohydrates (O’Shannessy et al., 1984; Bayer et al., 1987; Hoffman and O’Shannessy, 1988; O’Shannessy and Wilchek, 1990). Limited oxidation of carbohydrates or glycans using sodium periodate can create aldehydes on either sialic acid groups alone or on any sugar of the glycan pos-sessing adjacent hydroxyl groups (diols) (see Chapter 2, Section 2). Oxidized glycoproteins then may be reacted with a hydrazide-containing support to immobilize the proteins through hydrazone linkages, which only target the glycan portions of the molecules. This site-directed coupling method through the carbohydrate portion often aids in preserving active sites or binding regions within proteins. This can lead to better preser-vation of enzyme activity or better binding performance to a target molecule than using more random coupling methods like amine-reactive chemistries, which attach through lysine or N-terminal amines in many locations across the protein surface (Domen et al., 1990).

Glycosylated antibodies, for instance, can be immo-bilized onto a hydrazide-containing support after mild periodate oxidation of the carbohydrate, which is usu-ally located between the heavy chains within the Fc region. (Note: Some carbohydrate may be present on the Fab fragments near the antigen binding area in cer-tain antibody types.) The use of hydrazone-mediated coupling might be used to avoid an attachment of the antibody to the support near the antigen binding regions. The kinetics of periodate oxidation with anti-bodies to generate aldehydes was investigated by Hage et al. (1997) and found to involve two distinct popula-tions of oxidizable groups—one group that oxidized quickly within 5 to 10 min (likely sialic acid residues) and another group that took several hours to com-pletely oxidize. The oxidation of rabbit polyclonal IgG with 10-mM periodate resulted in the production of approximately one aldehyde per immunoglobulin mol-ecule after 10 min at 25°C, two aldehydes after 30 min, and about three aldehydes after 60 min. The rate of periodate oxidation of antibody carbohydrate also was found to be dependent on the pH of the reaction. From pH 3 to 7, the oxidation was found to decrease as the pH increased (Wolfe and Hage, 1995). After 30 min of oxidation at room temperature, two aldehydes were formed at pH 5 to 6, three were created at pH 4, and six resulted from the reaction at pH 3. Over-oxidation by extending the length of the periodate incubation proce-dure or by using concentrations of periodate exceeding



50-mM should be avoided to prevent damage to the avidity or immunoreactivity of antibodies (Abraham et al., 1991). Normally, periodate oxidation reactions are not done for longer than about 15 to 30 min, depend-ing on the sensitivity of the protein being treated. Treatment of antibodies with periodate at low concen-trations for 30 min is thus sufficient to provide alde-hydes on each antibody for efficient coupling while avoiding the damaging effects of over-oxidation.

Small-molecule aldehydes or ketones may also react with a hydrazide-activated support and end up being immobilized through hydrazone linkages. The reduc-ing ends of sugars, carbohydrates, or even glycans that have been cleaved off of glycoproteins can be selec-tively immobilized through this process. The reducing ends of carbohydrates typically react with the hydra-zide support at a slower rate due to the small amount of time most reducing sugars are in the open aldehyde form in solution as opposed to the cyclic hemiacetal form. Some biological small aldehyde- or ketone-con-taining molecules that may be present in some sam-ples such as pyridoxal, glyceraldehyde, acetaldehyde, pyruvate, α-ketoglutarate, acetone, and retinal can also couple to the support quite effectively and block some of the hydrazides to further coupling to a glycoprotein. For this reason, exposure of a hydrazide-containing matrix to crude biological samples should be avoided. In addition, never wash a hydrazide support with ace-tone or any other organic solvent containing a ketone, because the support will become inactivated.

Hydrazide-containing supports can be prepared by the coupling of a bis-hydrazide compound to a matrix using at least two reaction strategies. A bis-hydrazide such as adipic dihydrazide or carbohydrazide can be added in large excess to the appropriately prepared matrix to form spacer arms, which then terminate in free hydrazide groups. This process can be done through direct coupling to an amine-reactive support or through the intermediary use of a carboxylate-terminal spacer arm to which the bis-hydrazide compound is then coupled using EDC. Early methods described for

the preparation of a hydrazide-support involved the reaction of adipic acid dihydrazide with a periodate-oxidized agarose matrix (Hoffman and O’Shannessy, 1988). However, the reactivity of this aldehyde-con-taining support is so great toward the bis-hydrazide spacer that a large portion of the gel ends up being tre-mendously crosslinked, even to the extent of forming particle aggregates and damaged beads. This occurs even when adipic dihydrazide is added to the support in large mole excess, indicating that the potential for hydrazone formation is so high that both ends of the bis-hydrazide compound inevitably end up reacting with aldehydes on the support.

A more controlled approach to making a hydra-zide-activated support is to build it from a terminal carboxylate spacer arm derivative, which is coupled to the matrix initially and then activated to attach the bis-hydrazide (see section on spacer arms, this chap-ter). Activated carboxylates will react with a hydra-zide group on a bis-hydrazide compound to form a secondary amide (hydrazino) bond with a short spacer arm, which then terminates in a free hydrazide group (Figure 15.84). The reaction rate of coupling the hydra-zide to the carboxylate is much less than it is with an aldehyde-containing support, so the matrix does not become highly crosslinked and damaged during the process. The reaction can be made to occur through activation of the carboxylate spacer using a carbodi-imide such as EDC or through the use of an NHS ester-activated support (see previous discussion on amine reactive activation methods, this chapter). NHS may also be added to an EDC reaction to generate an intermediate NHS ester and enhance the reaction rate with the bis-hydrazide molecules. The result of these reactions is the preparation of a hydrazide-activated support that is ready to covalently couple to aldehyde-containing ligands.

Hydrazide-activated supports are relatively stable in aqueous solution if a preservative is added to prevent growth. The coupling of carbonyl-containing ligands to the matrix can be done over a broad pH range, but

Carboxylate-containingsupport

+ Hydrazide-activated support

EDC

O

O

O

OHN

H2N NH

NH2

Adipic dihydrazide

O

OHN

NH

NH

NH2

O

FIGURE 15.84 Preparation of a hydrazide support from a carboxylate spacer using EDC.



is optimal under slightly acidic conditions (pH 5–6). It has also been found that the yield of hydrazone for-mation and the rate of the reaction can be increased dramatically by use of an aniline-catalyzed reaction process (Cordes and Jencks, 1962; Dirksen et al., 2006a,b; Dirksen and Dawson, 2008; Byeon et al., 2010). Aniline will react with an aldehyde group on the ligand to form an intermediate Schiff base between its aryl nitro-gen and the carbon atom of the carbonyl group of the aldehyde. The aniline imine intermediate more rapidly gets protonated in solution than the initial oxygen of the aldehyde or ketone carbonyl, and it is this species that is the ideal form which can react with the hydra-zide groups (Kohler, 2009). This aniline imine interme-diate is then rapidly attacked by the hydrazides on the support, which results in displacement of aniline with associated hydrazone bond formation with the hydra-zides (Figure 15.85). This two-step reaction process results in much faster hydrazone formation and ligand immobilization than reactions done without the aniline catalyst being present. Using aniline catalysis offers the potential of doing immobilization reactions under more neutral pH conditions while still maintaining high effi-ciency of hydrazone bond formation (Zeng et al., 2009).

HYDRAZIDE ACTIVATION PROTOCOL

The following protocol describes the modification of a carboxylate-containing support with a bis-hydrazide

compound to form a hydrazide-activated matrix. The prior creation of a carboxylate-containing support may be achieved by coupling a carboxylate-terminal spacer arm to a matrix using the methods described elsewhere in this chapter on spacer arm production. The bis- hydrazide compound used can be one of many; however, adipic dihydrazide probably is the most common. An alternative small bis-hydrazide reagent is carbohydrazide, which is just a single carbonyl group with two hydrazines. It is the most reactive dihydrazide available and can be used to minimize the amount of alkyl chains used in a spacer arm, especially to limit the potential for creating sites of hydro-phobic binding and nonspecificity in the final matrix.

1. Wash 100 ml of a carboxylate-containing support material with water (at least 3 bed volumes) to remove storage buffers and preservatives. Wash the gel with an additional 2 bed volumes of 0.1-M MES, pH 4.75 (reaction buffer). The washing steps may be carried out using a sintered glass filter suspended in a suction filter flask and applying a gentle vacuum to facilitate drawing the wash solutions through the support. Drain the support to a moist cake.

2. Prepare 100 ml of a solution of 0.5-M adipic dihydrazide in the reaction buffer. Alternative bis-hydrazide compounds may be used at the same concentration with success. Readjust the pH if necessary.

Hydrazide-activated support

O

OHN

NH

NH

NH2

O

O

OHN

NH

NH

O

H

O

Aldehyde-containing

ligand

NH2

Aniline

+

H

N

IntermediateSchiff base

H

N

NH2

Ligand coupling throughhydrazone linkage

FIGURE 15.85 Immobilization of an aldehyde-containing ligand using an aniline catalyst.



3. Add the washed support to the dihydrazide solution with mixing. Continue to stir the support slurry by using an overhead paddle stirrer while adding 3 g of the carbodiimide EDC (fresh).

4. React for 3 h at room temperature with constant stirring.

5. Wash the support extensively with water, 1-M NaCl, and again with water to remove unreacted dihydrazide and any reaction byproducts. Store the hydrazide-containing support until use at 4°C as a 50% slurry in water containing a preservative.

LIGAND COUPLING TO HYDRAZIDE-ACTIVATED SUPPORTS

The following protocols describe the periodate oxi-dation of an antibody followed by its aniline-catalyzed immobilization onto a hydrazide-containing support. A subsequent immobilization protocol describes the immobilization of sugars, polysaccharides, or glycans through their reducing ends. Similar protocols can be used to immobilize other glycoconjugates or carbo-hydrates after periodate oxidation. In addition, small ligands containing aldehydes may be coupled to a hydrazide-containing support using the identical buf-fers and aniline addition process. Larger quantities of immobilized ligands may be prepared by proportion-ally increasing the amount of reagents used in these procedures.

(A) Periodate Oxidation of Antibody or Glycoprotein1. Dissolve 1 to 10 mg of an IgG antibody that is

glycosylated (or another glycoprotein) in 1 ml of coupling buffer (0.1-M sodium acetate, 0.15-M NaCl, pH 5.5). Be careful that the antibody does not contain an amine-containing buffer in solution, such as glycine or Tris, which may alter the pH of the coupling buffer or interfere with the subsequent coupling reaction. Dialyze or desalt the antibody using coupling buffer if necessary to remove interfering components before proceeding.

2. Weigh out 2.1 mg of sodium periodate into a small centrifuge tube and add the antibody solution to it with mixing using a vortex mixer. Continue to mix until the sodium periodate is completely dissolved and then wrap the tube in aluminum foil to protect it from light.

3. React for 30 min at room temperature with periodic mixing. Do not allow the oxidation to continue longer than this time period or oxidative damage may occur to the protein structure.

4. Stop the reaction by desalting the antibody solution using a size exclusion chromatography support having a molecular weight exclusion limit of no more than 10 kDa (i.e., a 10-ml column of Sephadex G-25 or

the equivalent). Use coupling buffer to perform the chromatography and collect the protein peak, which will elute in the void volume before the salt peak. Spin columns may also be used for this operation, as they will result in less dilution of the protein solution during the separation and are quicker to use (e.g., Zeba Spin Desalting Columns from Thermo Fisher). Use the oxidized antibody or glycoprotein in the coupling reaction immediately to prevent the potential for protein crosslinking over time through Schiff base formation or Mannich reaction processes.

(B) Coupling an Oxidized Antibody to a Hydrazide Support1. Wash 1 ml of a hydrazide-containing support with

water to remove storage solutions and preservatives. Then wash with several ml of coupling buffer (0.1-M sodium acetate, 0.15-M NaCl, pH 5.5). The washing of a small quantity of a chromatography support may be done in a drip column or a spin column without a top frit. Drain the gel to a wet cake and place the bottom cap on the column to stop the flow. The coupling reaction may be done in a sealed column or the washed gel transferred to a small plastic centrifuge tube able to hold at least 2.5 ml of slurry.

2. Add the oxidized antibody or glycoprotein from part (A) to the washed gel cake and stir to resuspend the matrix.

3. In a fume hood, add 18 μl of aniline catalyst to the gel slurry with stirring. This results in approximately a 0.1-M aniline solution in the coupling reaction mixture.

4. Continue the reaction for at least 2 to 4 h with constant mixing. Proteins with higher glycosylation content will couple faster than proteins with lower amounts of carbohydrate. The mixing may be done by end-over-end rocking in a sealed container or column.

5. Blocking of excess hydrazide groups on the support may be done by the addition of glyceraldehyde to the reaction mixture at a final concentration of 0.1-M and continuing to mix for 30 min. In most cases of glycoprotein immobilization, blocking of the unreacted hydrazides is not necessary; however, if this step is performed, do not add any reducing agents to the mixture or glyceraldehyde-protein adducts also will be formed (Acharya and Manning, 1980).

6. Wash the support with several bed volumes of coupling buffer and collect the washes. The amount of protein coupled may be determined by the difference in the amount of protein present in the reaction medium before coupling and after coupling, taking into account volume differences. Note that aniline may interfere with some methods of protein



concentration determination (e.g., absorbance at 280 nm) and may have to be removed. Continue to wash the support extensively with water, 1-M NaCl, and water to remove the last traces of all reactants.

7. The hydrazone linkages on the support may be reduced to stabilize them, if necessary. Often with coupling glycoproteins or glycosylated antibodies, the reduction step is not needed, because multi-point attachment takes place to the matrix and creates a high avidity bond. However, if reduction is deemed necessary due to a slow leakage of ligand from the support, then in a fume hood add to the washed gel 1 ml of coupling buffer containing 100-mM sodium cyanoborohydride (6.2 mg) (dangerous compound!). Reduce the hydrazone bonds for 1 h with constant mixing. Finally, wash the support as in step (6) and store the affinity support as a 50% slurry in water or buffer containing a preservative at 4°C.

(c) Coupling the Reducing end of Carbohydrates to Hydrazide Supports

Many carbohydrates, including monosaccharides, disaccharides, oligosaccharides, and glycans, contain a reducing sugar at their anomeric end that is in a cyclic hemiacetal form containing a masked aldehyde group. The aldehyde is only available for reaction when it is in

the open form, which is a minority of time in aqueous solution, so immobilization reactions with hydrazide-containing supports usually take much longer to go to completion than reactions with freely available alde-hydes. The coupling of reducing sugars onto hydrazide supports can be achieved with or without the pres-ence of a reducing agent and the result will be two dif-ferent linkage types (Figure 15.86). If a reductant such as sodium cyanoborohydride is used in the reaction medium, then coupling will take place at the anomeric carbon atom through ring opening and the formation of a reduced hydrazone linkage. However, if the reaction is performed without a reducing agent being present, the result will be the creation of a glycosylhydrazide bond with an intact ring structure at the reducing sugar (Rothenberg et al., 1993; Toomre and Varki, 1994; Bigge et al., 1995; Leteux et al., 1998; Srikrishna et al., 2001). It may be important for some affinity chromatography applications to maintain the conformation of a cyclic sugar structure at the reducing end in order to ensure interaction with certain lectins.

The addition of an aniline catalyst to the coupling reaction with a reducing sugar has been shown to dra-matically accelerate the formation of the hydrazone bond (Thygesen et al., 2010). The aniline initially traps the open ring form of the sugar’s aldehyde through


NH2NH

O

+ OO

OHHO

HO

OH

Beta-D-glucosewith accessiblereducing end

O

OOHHO

OH

HOO

OHHO

O

OHAldehydeHemiacetal

NaCNBH3

No reductant

NH

O

HN

NH

O

HO

OOHHO

HN

OH

Glycosylhydrazide bond

Secondary amine linkage

FIGURE 15.86 Coupling at the reducing end of a carbohydrate with and without a reducing agent.



the formation of an imine, which then gets attacked by the hydrazide to form the desired hydrazone with the matrix (Figure 15.87). The following protocol describes the catalyzed reaction with aniline being present in the coupling buffer.

1. Dissolve a carbohydrate, polysaccharide, or glycan (having a reducing end available) at a concentration of at least 5 to 100 μM in 1 ml of 0.1-M sodium acetate, 0.15 M NaCl, pH 5.5. Note: Higher concentrations may be used for more abundant carbohydrates as appropriate. The optimal concentration of the carbohydrate may have to be determined experimentally by making several small batches and varying the amount of ligand added to the reaction. Ultimately, the best performance of the immobilized carbohydrate in the intended affinity application should be used as the determining factor for the best concentration to use in the coupling reaction. An alternative coupling medium is a nonaqueous environment consisting of 30% glacial acetic acid in DMSO (v/v) or an acetic acid/pyridine mixture of 1 : 2 (v/v). Nonaqueous conditions may facilitate the dissolution of some carbohydrate molecules better than aqueous buffers.

2. Wash 1 ml of a hydrazide-containing support with water to remove storage solutions and preservatives. Then wash with several milliliters of coupling buffer

(0.1-M sodium acetate, 0.15-M NaCl, pH 5.5). The washing of a small quantity of a chromatography support may be done in a drip column or a spin column without a top frit. Drain the gel to a wet cake and place the bottom cap on the column to stop the flow. Note: If an organic solvent medium is to be used for the immobilization reaction, sequentially wash the matrix into the acetic acid/solvent blend by first washing it into 100% solvent without the acid to completely remove water (e.g., DMSO) and then washing with the desired acetic acid/solvent mixture (e.g., 30% acetic acid/DMSO).

3. Add the carbohydrate solution to the washed gel cake and stir to resuspend the matrix. Transfer the slurry to a centrifuge tube that is large enough to hold a total volume of at least 4 ml.

4. In a fume hood, add 18 μl of aniline catalyst to the 2 ml gel slurry with stirring. This results in approximately a 0.1-M aniline solution in the coupling reaction mixture.

5. Seal the tube and continue the reaction for at least 4 h at room temperature and with constant mixing by end-over-end rocking.

6. The blocking of excess hydrazide groups on the support may be achieved by the addition of glyceraldehyde to the reaction mixture at a final concentration of 0.1-M and continuing to mix for 30 min.


NH2NH

O

OO

OHHO

HO

OH

Beta-D-glucosewith accessiblereducing end

HOO

OHHO

O


HOO

OHHO

OH

Aniline N

Intermediate anilineSchiff base

Ring open form

Carbohydrate coupled viahydrazone linkage

NH

O HO

OOHHO

OH

N

NH2

FIGURE 15.87 Immobilization through the reducing end of a carbohydrate using aniline.



7. The hydrazone linkages on the support may be reduced to stabilize them, if necessary. Often when coupling through a single hydrazone bond to a ligand, as in the reaction with the reducing end of a carbohydrate or glycan, the reduction step can be performed to prevent a slow leakage of ligand from the support. To the washed gel add 1 ml of coupling buffer containing 100-mM sodium cyanoborohydride (6.3 mg) (dangerous compound—use a fume hood!). Reduce the hydrazone bonds for 1 h with constant mixing.

8. In the fume hood, wash the support extensively with water, 1-M NaCl, and water to remove the last traces of all reactants and store the affinity support as a 50% slurry in water or buffer containing a preservative at 4°C.

Aminooxy Supports for Coupling Aldehydes or Ketones

The chemoselective ligation reaction of an aldehyde group with an aminooxy group (–ONH2) to yield an oxime bond (aldoxime) has been described for use in many bioconjugation reactions, as well as in the coupling of ligands to insoluble supports including surfaces, for the organic synthesis of radiohalogenated compounds, and in the synthesis of drug candidates (Thumshirn et al., 2003; Poethko et al., 2004; Liu et al., 2007; Colombo and Bianchi, 2010). This reaction with aminooxy groups also is quite efficient with ketones to form an oxime called a ketoxime. The simplest form of this reaction has been known for over a century and occurs with alde-hydes or ketones as they react with the small compound hydroxylamine (Figure 15.88). The conversion of cyclo-hexanone to its oxime with hydroxylamine followed by subsequent treatment with sulfuric acid to yield capro-lactam via a Beckmann rearrangement produces a raw material used in billions of kilogram quantities to create nylon-6 polymers by ring-opening polymerization.

Hydroxylamine derivatives that consist of an organic group linked to the oxygen also are very effective at

forming oxime bonds with carbonyl groups. In this case, the hydroxylamine derivative typically is called an aminooxy functional group, which is also known as an aminoxy or alkoxyamine derivative. The aminooxy compound of this reaction contains a substituent off the oxygen group of hydroxylamine, which can consist of virtually any organic compound or even a chromatog-raphy support or modified surface containing a spacer arm that terminates in the –ONH2 group. The immo-bilization of aldehyde- or ketone-containing affinity ligands onto support materials that have been activated to contain aminooxy groups can be done similar to the methods described previously for hydrazide contain-ing supports. However, oxime formation usually occurs more rapidly than the reaction between a hydrazide group and an aldehyde or ketone and it typically pro-vides more stable oxime bonds than hydrazones.

The formation of an aminooxy group on a chroma-tography support can be achieved through the creation of the appropriate spacer arm. There are many meth-ods that can be used to build a spacer on a support (see section on spacer arms, this chapter). In most cases, the creation of the spacer must be performed using a pro-tected aminooxy group, because the terminal amino group will react in amine coupling reactions, which are typically performed to build the spacer molecules. For instance, the first step in making a support with amino-oxy functionalities might be to couple a diamine spacer to an amine-reactive support (see section on amine reac-tive immobilization methods, described previously in this chapter). This would provide an amine-containing intermediate that could then be used to link a protected aminooxy-containing molecule, such as phthalimidooxy –dPEG12–NHS ester (Quanta Biodesign). This crosslinker contains a phthalimide-protected aminooxy group on one end and an amine-reactive NHS ester on the other end which can be used to modify the amine-containing support. The result of reacting with the NHS ester ends will be the formation of amide bonds with the amine groups on the support, which will create extremely hydrophilic PEG12 chains that extend off the matrix and terminate in the protected aminooxy functional-ities. Subsequent removal of the phthalimide-protecting groups with hydrazine in aqueous solution yields the free aminooxy-activated support (Figure 15.89).

Another approach to forming aminooxy functional-ities on a resin is to make use of an activated support containing tosyl groups (see previous section on tosyl activation, this chapter). This reactive group is usually used to immobilize amine-containing ligands, but it can also be used to react with hydroxyl groups in nonaque-ous conditions. The reaction of a tosyl-activated sup-port with the compound N-hydroxyphthalimide results in coupling through the available hydroxyl group on the reagent. This intermediate will result in a protected

R O

H

R1 R2

O

Aldehydecompound

Ketonecompound

+

+

H2N OH

Hydroxylamine

R NOH

H

R1 NOH

R2

Aldoximelinkage

Ketoximelinkage

H2N OH

Hydroxylamine

FIGURE 15.88 Basic reaction of an aldehyde or ketone with ami-nooxy group to give an oxime bond.



aminooxy group immobilized on the support, which then can be deprotected by the addition of 0.5-M hydra-zine in aqueous solution, thus producing the aminooxy-activated matrix (Figure 15.90).

Aminooxy groups have also been used on planar substrates to covalently array proteins to form nanopat-terns on silicon wafers and other surface materials. In one example, the aminooxy functionalities were coated on the surface of wafers by Christman et al. (2008) using eight-arm PEG derivatives. The hydroxyl groups on the outer ends of the PEG arms were modified using a Mitsunobu-type reaction with N-hydroxyphthalimide and including triphenyl phosphine (PPh3) with DIAD (diisopropyl azodicarboxylate) as the activators (Figure 15.91). Subsequent deprotection and removal of the phthalimide using hydrazine yielded the desired aminooxy groups.

The immobilization reaction using an aminooxy sup-port with aldehyde- or ketone-containing ligands can be accelerated by the addition of an aniline catalyst. As in the formation of hydrazone linkages with aldehydes and hydrazide-activated supports, the formation of

an oxime bond between an aldehyde or ketone and an aminoxy is dramatically increased in rate and yield by aniline (Cordes and Jencks, 1962; Dirksen et al., 2006a,b; Dirksen and Dawson, 2008; Byeon et al., 2010). The aro-matic amine on aniline first reacts with the aldehyde or ketone group to form an intermediate imine, which then reacts with the aminooxy groups on the support to create the final oxime linkage (Figure 15.92). The intermediate aryl imine undergoes more rapid proton-ation in slightly acidic solution than does the oxygen of the aldehyde or ketone carbonyl, and it is this proton-ated species that can react with the aminooxy groups (Kohler, 2009).

The immobilization reaction on a surface that was activated with aminooxy groups to couple a peptide molecule was studied in detail to determine its kinet-ics (Lempens et al., 2009). The peptide was prepared to contain an N-terminal aldehyde residue made via the pyridoxyl 5′-phosphate/sodium periodate method of Gilmore et al. (2006a,b) (also see Scheck and Francis, 2007; Scheck et al., 2008). It was also found in this case that use of the aniline catalyst dramatically increased

O

HN

NH2


primary amines

+ N

O

O

OO O

ON

O

O12

O

HN

NH

Phthalimidooxy–dPEG12–NHS ester

O

OO

N

O

O12

0.5 Mhydrazine

O

HN

NH

O

OO

NH212

+HN

HN

O

OAminooxy–PEG12 support

FIGURE 15.89 Preparation of an aminooxy support using PhthNO–dPEG12–NHS ester.



the reaction rate and coupling efficiency at all pH values investigated, with the greatest rate and yield obtained at pH 4.5. The rate at pH 6.0 was less than half that observed at the lower pH value; however, the dif-ference between a catalyzed and an uncatalyzed reac-tion was significant at all the acidic pH environments studied. As the reaction pH is increased from 4.5 to more of a neutral pH environment, the rate and yield of coupling significantly slow down; however, for mol-ecules sensitive to more acidic pH environments the reaction may be performed with good results at higher pH. To compensate for slower kinetics at higher pH, the time of the reaction should be increased to realize acceptable immobilization yields.

AMINOOXY ACTIVATION PROTOCOL

The following protocol describes the modification of an amine-reactive support with a diamine spacer and the subsequent reaction of the amino-support with an NHS–PEG12–oxyphthalimide crosslinker followed by deprotection of the phthalimide to form the amino-oxy derivative. Other amine-containing supports may also be used in this procedure, such as can be prepared using the spacer arms described elsewhere in this chap-ter. Use a fume hood and personal protective equip-ment for all operations.

1. In a fume hood, drain 10 ml of a CDI-activated agarose support of acetone storage solution (see section on CDI activation). Other amine-reactive supports may also be used to couple the diamine; however, follow the respective protocols for coupling ligands if another reaction is used. All of the washing steps may be carried out using a sintered glass filter suspended in a suction filter flask and applying a slight vacuum to facilitate drawing the wash solutions through the support. Pull a gentle vacuum on the suction filter flask and break up the resin into small pieces as the acetone drains through, but do not allow the support to dry. Next, add to the resin approximately 2 bed volumes of the solvent DMAC (note that DMSO may be used as an alternative solvent) and stir the gel to resuspend it in the new solvent. Continue to wash the support with DMAC for at least 5 bed volumes to remove most of the remaining acetone.

2. Prepare 10 ml of a solution consisting of 1.0-M ethylenediamine in DMAC (or DMSO). Notes: Alternative diamine compounds may be used at the

O


SO

OCH3 +

N

O

O

HO

N-hydroxyphthalimide

N

O

O

O

O SO

OCH3

0.5 M hydrazine

+HN

HN

O

O

NH2O

Aminooxy support

FIGURE 15.90 Formation of aminooxy groups on a support by reaction of N-hydroxyphthalimide with a tosyl-activated support.

NN

O

OO

O

DIAD:diisopropyl azodicarboxylate

OH


+N

O

O

HO

N-hydroxyphthalimide

Triphenylphosphine

(PPh3)

CH2Cl2

N

O

O

O

0.5 Mhydrazine

+HN

HN

O

O

NH2O

Aminooxy support

Mitsunobureaction

FIGURE 15.91 Aminooxy formation at the end of a PEG chain using the Mitsunobu reaction.



same concentration with success; however, avoid the use of long aliphatic diamines as spacers, as this will create considerable hydrophobic character within the matrix and increase the potential for nonspecific interactions on the subsequent affinity support.

3. Add the washed support to the ethylenediamine solution with stirring. React for 1 h at room temperature with constant mixing. End-over-end rocking in a sealed container may be used to mix the support. Note: The addition of an organic base to the reaction to accept protons generated during the coupling of the spacer need not be done, because the free amine end of ethylenediamine will still be available.

4. Wash the amine-containing support extensively with DMAC to remove the last traces of diamine compound (use at least 20 bed volumes). Drain the washed support to a moist cake.

5. Prepare a solution of the heterobifunctional crosslinker PhthNO–dPEG12–NHS ester (Quanta Biodesign) in 10 ml of DMAC dissolved at a concentration of at least 10 mg/ml (11.6 μmol/ml). Higher concentrations may be used to create higher densities of the final aminooxy groups on the matrix.

6. Add the washed support to the crosslinker solution and mix to resuspend the gel. To this suspension, add 0.65 g of dimethylaminopyridine (DMAP) as an organic base to catalyze the reaction and

mix to dissolve (alternatively, the addition of 0.75 ml of anhydrous triethylamine (TEA) may be done or an equivalent mole amount of N,N-diisopropylethylamine (DIEA) may be added). React for 1 h at room temperature with constant mixing.

7. Extensively wash the modified support with DMAC to remove excess reactants and reaction byproducts (at least 20 bed volumes). Next, wash the support into water by sequentially washing with increasing concentrations of water in DMAC until 100% water is used. Continue washing with water until all traces of the solvent have been removed. Drain the gel to a wet cake.

8. In a fume hood, prepare 10 ml of an aqueous solution consisting of 0.5-M hydrazine (dangerous and toxic!) in 0.1-M MES, pH 6.0. If hydrazine dihydrochloride is used to prepare this solution, its dissolution in the buffer will not cause the pH to become highly alkaline. Avoid contact with hydrazine or its solutions by using the appropriate personal protective equipment, including being cautious to avoid contact with skin or eyes, and also avoiding the inhalation of dust. Prevent static charge buildup when dispensing and weighing the hydrazine compound by using the appropriate grounding precautions. Adjust the final pH of the solution back to 6.0, if necessary.

H

O

Aldehydecontaining

ligand

NH2

Aniline

+

H

N


NH2

Ligand coupling throughoxime linkage

NH2O

Aminooxy support

O

H

N

FIGURE 15.92 Immobilization of an aldehyde ligand on an aminooxy support using aniline catalysis.



9. In the fume hood, add the washed gel containing the protected aminooxy groups with the hydrazine solution and mix by end-over-end rotation in a sealed container. React overnight to deprotect the aminooxy groups.

10. In the fume hood, drain the support of excess hydrazine deprotection reagent and wash the support extensively with water, 1-M NaCl, and water. Store the aminooxy-activated support until use as a 50% slurry in water or buffer containing a preservative at 4°C. (Note: Do not include any ketone- or aldehyde-containing compounds in the storage solution or the aminooxy groups will react and become inactive for coupling ligands.)

LIGAND COUPLING TO AMINOOXY-ACTIVATED SUPPORTS

The following protocols describe the periodate oxi-dation of sialic acid groups on glycoproteins followed by their aniline-catalyzed immobilization onto an ami-nooxy-containing support. A subsequent immobiliza-tion protocol describes the immobilization of sugars, polysaccharides, or glycans through their reducing ends. Similar protocols can be used to immobilize other glycoconjugates or carbohydrates after periodate oxi-dation or through their reducing ends, if available. In addition, small ligands containing aldehydes or ketones may also be coupled to an aminooxy-containing sup-port using the identical buffers and an aniline catalysis process. Larger quantities of immobilized ligands may be prepared by proportionally increasing the reagent amounts used in these procedures.

(A) Periodate Oxidation of Sialic Acid Groups on Glycoproteins1. Dissolve 1 to 10 mg of a glycoprotein containing sialic

acid groups in 1 ml of coupling buffer (0.1-M sodium acetate, 0.15-M NaCl, pH 5.5). Be careful that the glycoprotein does not contain an amine-containing buffer in solution, such as glycine or Tris, which may alter the pH of the coupling buffer or interfere with the subsequent coupling reaction. For instance, dialyze or desalt a commercial antibody preparation containing Tris or glycine using coupling buffer to remove interfering components before proceeding. Chill the glycoprotein solution by placing it on ice.

2. Dissolve sodium periodate in water at a concentration of 10 mg/ml (46-mM). Continue to mix until the sodium periodate is completely dissolved and then wrap the tube in aluminum foil to protect it from light. Chill the periodate solution by placing it on ice.

3. Add 21.8 μl of the periodate solution to the 1 ml of glycoprotein solution and mix to dissolve (makes approximately 1-mM periodate final concentration in the glycoprotein solution). Maintain the solution on ice.

4. React for 30 min on ice with periodic mixing. Do not allow the oxidation to continue longer than this time or oxidation may occur at sites other than just sialic acid groups.

5. Stop the reaction by desalting the antibody solution using a size exclusion chromatography support having a molecular weight exclusion limit of no more than 10 kDa (i.e., at least 5 ml of Sephadex G-25 or the equivalent). Use cold coupling buffer to perform the chromatography and collect the protein peak, which will elute before the salt peak. Spin columns may also be used for this operation, as they will result in less dilution of the protein solution during the separation and are quicker to use (e.g., Zeba Spin Desalting Columns from Thermo Fisher). Use the oxidized glycoprotein in the coupling reaction immediately to prevent the potential for protein crosslinking over time through Schiff base formation or Mannich reaction processes.

(B) Coupling the Oxidized Glycoprotein to an Aminooxy Support1. Wash 1 ml of an aminooxy-containing support with

water to remove storage solutions and preservatives. Then wash with several milliliters of coupling buffer (0.1-M sodium acetate, 0.15-M NaCl, pH 5.5). The washing of a small quantity of a chromatography support may be done in a drip column or a spin column without a top frit. Drain the gel to a wet cake and place the bottom cap on the column to stop the flow. The coupling reaction may be done in a sealed column or the washed gel transferred to a small plastic centrifuge tube able to hold at least 2.5 ml of slurry. Note: The pH of the coupling buffer may be decreased to pH 4.5 to further enhance the reaction rate and yield of oxime bond formation.

2. Add the oxidized glycoprotein from part (A) to the washed gel cake and stir to resuspend.

3. In a fume hood, add 18 μl of aniline catalyst to the gel slurry with stirring. This results in approximately a 0.1-M aniline solution in the coupling reaction mixture.

4. Continue the reaction for at least 2 to 4 h with constant mixing. Proteins with higher glycosylation content will couple faster than proteins with lower amounts of carbohydrate. The mixing may be done by end-over-end rocking in a sealed container or column.

5. Blocking excess aminooxy groups on the support may be done by the addition of glyceraldehyde to the reaction mixture at a final concentration of 0.1-M and continuing to mix for 30 min. In most cases of glycoprotein immobilization, blocking the unreacted aminooxy groups is not necessary; however, if this step is performed, do not add any reducing agents to the mixture or glyceraldehyde–protein adducts will be formed (Acharya and Manning, 1980).



6. Wash the support with several bed volumes of coupling buffer and collect the washes. The amount of protein coupled to the resin may be determined by the difference in the amount of protein present in the reaction medium before coupling and that remaining after coupling, taking into account volume differences. Note that aniline may interfere with some methods of protein concentration determination (e.g., absorbance at 280 nm) and may have to be removed. Continue to wash the support extensively with water, 1-M NaCl, and water to remove the last traces of all reactants. Finally, wash the support as in step 6 and store the affinity support as a 50% slurry in water or buffer containing a preservative at 4°C.

(C) Coupling the Reducing end of Carbohydrates to Aminooxy Supports

Many carbohydrates contain a reducing sugar at their anomeric end that is in a cyclic hemiacetal form, which in reality is a masked aldehyde group. The alde-hyde only is available for immobilization when it is in

the open form, which is a minority of time in aqueous solution. For this reason, the immobilization reactions of reducing sugars and carbohydrates with aminooxy-containing supports usually take longer to go to com-pletion than reactions with freely available aldehydes. The coupling of reducing sugars onto aminooxy sup-ports is carried out very similarly to that described for hydrazide supports, described previously; however, a reducing step is optional to further stabilize the resul-tant bond. If the reaction is carried out without a reduc-ing agent being present, the result will be the creation of an oxime bond yielding a mixture of an intact ring structure (glycosyl derivative) or acyclic oxime (open ring derivative) at the reducing sugar anomeric car-bon. If a reductant is present during the reaction, only the acyclic derivative will result with the formation of a glycosyl-hydroxylamine linkage to the support (Peluso et al., 2002) (Figure 15.93).

The addition of an aniline catalyst to the coupling reaction of an aminooxy group with a reducing sugar has been shown to dramatically accelerate the for-mation of the oxime bond (Thygesen et al., 2010). The

OO

OHHO

HO

OH

Carbohydratewith availablereducing end

HOO

OHHO

O


O

HO

OOHHO

HN

OH

Acyclic secondaryaminooxy linkage

NH2O

Aminooxysupport

O

HO

OOHHO

N

OH

+

O

OOHHO

OHHN

O

Cyclic glycosylhydroxylamine bond

With reductant Without reductant

Acyclic oxime derivative

FIGURE 15.93 Immobilization of reducing carbohydrate onto an aminooxy support with and without the use of a reductant.



aniline initially traps the open ring aldehyde through the formation of an imine, which then gets attacked by the immobilized aminooxy group to form the desired oxime bond with the matrix (Figure 15.94). The fol-lowing protocol describes the catalyzed reaction with aniline being present in the coupling buffer. Larger quantities of the immobilized ligand may be prepared by proportionally increasing the amount of reagents added at each step.

1. Dissolve a carbohydrate, polysaccharide, or glycan (having a reducing end available) at a concentration of at least 5 to 100 μM in 1 ml of 0.1-M sodium acetate, 0.15-M NaCl, pH 4.5. Note: Higher concentrations may be used for more abundant carbohydrates as appropriate. The optimal concentration of the carbohydrate may have to be determined experimentally by making several small batches and varying the amount of ligand added to the reaction. Ultimately, the best performance of the immobilized carbohydrate in the intended affinity application should be the determining factor for the best coupling reaction conditions to use.

2. Wash 1 ml of an aminooxy-containing support with water to remove storage solutions and preservatives. Then wash with several milliliters of coupling buffer (0.1-M sodium acetate, 0.15-M NaCl, pH 4.5). The washing of a small quantity of a chromatography support may be done in a drip column or a spin column without a top frit. Drain the gel to a wet cake and place the bottom cap on the column to stop the flow.

3. Add the carbohydrate solution to the washed gel cake and stir to resuspend the matrix. Transfer the slurry to a centrifuge tube that is large enough to hold a total volume of at least 4 ml.

4. In a fume hood, add 18 μl of aniline catalyst to the 2-ml gel slurry with stirring. This results in approximately a 0.1-M aniline solution in the coupling reaction mixture.

5. Seal the tube and continue the reaction for at least 4 h at room temperature and with constant mixing by end-over-end rocking.

6. The blocking of excess aminooxy groups on the support may be done, if necessary, by the addition of glyceraldehyde to the reaction mixture at a final concentration of 0.1-M and continuing to mix for 30 min.

7. In the fume hood, wash the support extensively (at least 20 bed volumes) with water, 1-M NaCl, and water to remove the last traces of all reactants and store the affinity support as a 50% slurry in water or buffer containing a preservative at 4°C.

Amine-Containing Supports for Coupling Aldehydes, Ketones, or Carboxylates

Supports that have available primary amines on them can be used to immobilize carbonyl-contain-ing ligands with good efficiency. The amines may be formed from the construction of a spacer arm on a base support or created as a result of the polymerization of functional silane compounds or vinyl monomers con-taining amines. The amine groups can react with alde-hydes or ketones on affinity ligands using a reductive amination process to yield secondary amine linkages to the support. Free amine groups can also be used to immobilize carboxylate-containing ligands by amide bond formation using a carbodiimide-mediated reac-tion. The reaction of an immobilized amine with an aldehyde-containing ligand is just the opposite of the immobilization of an amine-containing ligand on an aldehyde support, which is described under the sec-tion Amine Reactive Immobilization Methods earlier in this chapter; however, the reaction principles are identical.

Amine-containing spacer arms can be added to a support by the reaction of a diamine compound with an amine-reactive matrix. Using this method, a variety

OO

OHHO

HO

OH

Carbohydratewith reducingend available

HOO

OHHO

O


NH2

Aniline

HOO

OHHO

OH

N

NH2O

Aminooxysupport

Aniline Schiffbase intermediate

NH2

O

HO

OOHHO

N

OH

Acyclic oxime derivative

FIGURE 15.94 Immobilization through the reducing end of a carbohydrate using aniline.



of spacer types may be used as desired to design par-ticular properties into the resin, such as the use of short or long hydrophobic or hydrophilic cross-bridges, which may have an effect on subsequent affinity sepa-rations done on the support. Amines also can be added to particles using a functional silane reagent, such as aminopropyltrimethoxysilane, that can be used to coat the surface of particles and form the required primary amine groups. See the final section of this chapter for a discussion on the formation of spacer arms on supports and see also Chapter 13 for methods related to the use of functional silanes.

An amine-containing support can react with ligands containing aldehydes or ketones to form an initial Schiff base imine, which results from a dehydration reaction. This imine bond is highly reversible in aque-ous environments and therefore must be stabilized by reduction to permanently link the ligand to the sup-port (Figure 15.95). The reducing agent used in this process typically is sodium cyanoborohydride or other equivalent reagents that are capable of reducing the imine to a secondary amine, but will not reduce the ini-tial aldehyde reactants. The use of an amine-containing support to immobilize aldehydes or ketones may not be the first or best choice, because hydrazide- or ami-nooxy-activated supports have better reaction charac-teristics and yield more stable linkages (see previous sections). However, if an amine-containing matrix is the desired starting point, the coupling of carbonyl ligands can be done with success using the cyanoborohydride reduction step to stabilize the linkage.

However, an amine-containing support is even more suitable for the immobilization of carboxylate-contain-ing ligands. Carboxylates and amines can be made to react and form stable amide bonds. Although these two functional groups do not spontaneously react under normal conditions, the addition of an amide bond form-ing agent can be done to facilitate efficient coupling. The most common amide bond forming agents are

carbodiimides, such as the water soluble EDC, as well as other such condensing agents that were originally developed to form bonds between amino acids in pep-tide synthesis applications. The water soluble reactants can be used in aqueous buffer to couple carboxylate-containing ligands that are soluble in water. EDC is a so-called “zero-length” crosslinker, since it mediates the formation of amide linkages without leaving behind a spacer molecule (Grabarek and Gergely, 1990). In addi-tion, for carboxylate ligands that may be insoluble in aqueous conditions, amide bond forming agents that are soluble in organic solvents can be used. EDC reacts with the carboxylates on the ligand to form an interme-diate reactive ester, which then goes on to react with the amines on the support surface to create amide bond linkages (Figure 15.96). See also Chapter 14 on the use of EDC for coupling of affinity ligands to microparticles and nanoparticles for additional information on the reactions involved with carbodiimide activation and immobilization.

In most cases, EDC-mediated amide bond forma-tion is quite efficient and proceeds to completion within 2 to 4 h. Staros et al. (1986) developed a modification of this reaction that incorporates the addition of NHS (or sulfo-NHS) into the reaction medium to form an inter-mediate NHS ester on the activated carboxylate. This two-stage reaction results in the creation of an amide bond through the reaction of an intermediate NHS ester with the amine groups on the support. An NHS ester undergoes fewer side reactions and is more efficient at forming amide bonds than the intermediate EDC ester, so the desired product is formed at an accelerated rate.

LIGAND COUPLING TO AMINE-CONTAINING SUPPORTS

The following protocols make use of an amine-con-taining resin made through the modification of an amine-reactive support with a diamine spacer molecule (see the last section in this chapter on spacer arm construction).

NH2


N+

H

O

Aldehyde-containingligand

H


HN

Ligand coupledvia secondaryamine bond

NaCNBH3

FIGURE 15.95 Immobilization of an aldehyde-containing ligand onto an amine-containing support.



Other amine-containing supports can also be used with success, such as polymeric supports prepared using amine-containing monomers during the polymeriza-tion process. Also see Chapter 14 for protocols related to the coupling of affinity molecules to micro- or nanoparticles.

Coupling Aldehydes or Ketones to Amine-Containing Supports

The following protocol makes use of sodium cyanobo-rohydride, which is a highly toxic compound that expels volatile cyanide into the atmosphere. For this reason, all operations using this compound as the solid or in solu-tion should be done in a well-ventilated fume hood. Also use the appropriate personal protective equipment while doing this procedure, such as gloves, safety glasses, and a lab coat to prevent direct contact with the compound.

1. Wash 10 ml of the amine-containing support material into coupling buffer (0.1-M sodium phosphate, pH 7.2) and drain to a wet cake. Other buffer components may be added to the coupling buffer, such as the addition of 0.15-M NaCl or alternative buffer salts; however, avoid amine-containing compounds like Tris, glycine, or imidazole, which will compete in the reaction. In general, avoid buffer compounds containing primary or secondary amines

as well as any other nucleophile that could react with an aldehyde on the ligand. In addition, certain detergents and other amphiphillic components have been shown to decrease the stability of the intermediate Schiff base and these additives should be used only after validating that they have no effect on the immobilization reaction (Viguera et al., 1990). Reductive amination coupling in aqueous solution has been shown to effectively occur between pH 4 and 10, with an optimal range for cyanoborohydride reduction of pH 6 to 8.

2. Suspend the washed support material containing amine residues in an equal volume of 0.1-M sodium phosphate, pH 7.2, into which an aldehyde-containing ligand has been dissolved. Notes: For glycoprotein ligands, a typical concentration range may be 3 to 5 mg/ml gel, but much higher concentrations may be used if a high density of coupled protein on the gel is desirable. In some cases, reactions containing protein at up to 20 mg/ml gel can be done, but the need for this density on the final affinity support is unusual. For small aldehyde-containing affinity ligands, a concentration of 2 to 3 mg ligand/ml gel in the immobilization reaction may be sufficient to obtain good binding capacity on the resultant affinity gel. Alternatively, a concentration representing 5 to 10 times the

H2N


+

OH

O

Carboxylate-containing

ligand

Intermediatereactive ester

EDC

NN

CNH

Cl

ON

NHC

HN Cl

O+

O

HN

NHC

HN Cl

O

NH

Immobilized ligandvia amide bond formation

Isoureabyproduct

FIGURE 15.96 Immobilization of carboxylate ligands onto amine-containing supports using EDC.



concentration of reactive amines on the matrix may be used to ensure a high density of the final coupled ligand. The ultimate concentration of ligand used in the reaction should be optimized by performing a series of coupling reactions at different initial ligand concentrations and determining the performance of the affinity supports in capturing and eluting the desired target molecule.

3. Stir the gel/ligand slurry in a fume hood using a paddle stirrer or using a rotator to maintain constant mixing.

4. Add to the gel slurry 63 mg of solid sodium cyanoborohydride (NaCNBH3; MW = 62.84) [toxic!] or 0.2 ml of 5-M NaCNBH3 in 1-N NaOH (Sigma) with stirring. If the alkaline solution of cyanoborohydride is used, check the pH and readjust to 7.2, if necessary. Continue the reaction for 4 h to overnight at room temperature. The amount of cyanoborohydride added—whether as the pure solid compound or as the solution—results in a 50-mM solution in the final reaction slurry, which is in a total volume of 200 ml. A shorter coupling time may be sufficient for many ligands, but the optimal conditions to give acceptable yields for a particular ligand should be determined by doing a series of coupling reactions at different time points. Reactions at 4°C may also be done for thermally sensitive proteins or ligands, but reaction times may have to be extended to get the same level of coupling as a room temperature reaction.


6. Unreacted amine residues remaining on the support can be blocked by the addition of a small aldehyde-containing compound, such as glyceraldehyde. Avoid the blocking of excess amine groups if a protein ligand or a ligand containing more than one amine has been immobilized, as the glyceraldehyde also will modify amines on the ligand. Wash the support once with an equal volume of 0.1-M glyceraldehyde dissolved in coupling buffer at pH 7.2, and then transfer the wet gel cake to a clean flask or vessel used for mixing the reaction. Add with stirring 10 ml of the 0.1-M glyceraldehyde solution along with 63 mg (or 0.2 ml of the 5-M NaCNBH3 solution) of sodium cyanoborohydride. Readjust the pH if necessary and continue the blocking reaction for 30 min at room temperature.

7. Wash the support extensively with water, 1-M NaCl, and again with water to remove all unreacted components from the gel. Additional wash solutions may be utilized to completely remove ligands that may have some nonspecific binding potential to remain noncovalently bound to the immobilized ligand, such as the use of acidic and alkaline washes

as well as washes containing denaturants (such as guanidine). After washing, the affinity gel may be stored as a 50% aqueous slurry containing a preservative at 4°C.

Coupling Carboxylate-Containing Ligands to Amine-Containing Supports

The following protocol may be used to immobilize a carboxylate-containing ligand onto a support material containing amines. The process uses the water soluble carbodiimide EDC, which activates the carboxylate to an intermediate ester that then reacts with the amines on the support to form amide bonds. This method is not recommended for coupling ligands that contain both carboxylates and amines, such as proteins, because the ligand will become oligomerized in solution upon the addition of EDC.

1. Wash 10 ml of an amine-containing support with several volumes of water to remove storage solutions and then wash with coupling buffer (0.1-M MES, pH 4.7). Drain to a wet cake. Note: The EDC reaction occurs efficiently at slightly acidic pH values with an optimal rate in the range of pH 4.5 to 6.0. However, buffers at physiological pH may also be used with success (e.g., 0.1-M sodium phosphate, pH 7.0–7.5). The reaction at the higher pH will be somewhat slower, and the reaction time should be extended to obtain the same yield of coupling. For neutral pH reactions, the addition of 2 equivalents of NHS (or sulfo-NHS) over the amount of EDC added can be done to further accelerate the reaction kinetics.

2. Prepare 10 ml of the carboxylate-containing ligand to be coupled in coupling buffer. For many small ligands, the use of 3 to 5 mg ligand/ml is a sufficient concentration; however, some optimization of ligand concentration may have to be done to obtain the best performance of the affinity resin in the intended application. Note: For ligands that are not very soluble in aqueous solution the carboxylate molecule may be first dissolved in a water-miscible solvent such as ethanol, DMSO, DMF, or DMAC and then added to the coupling buffer to obtain the final mixture. The final percentage of solvent in the coupling buffer should not exceed about 20% for DMSO, DMF, or DMAC or the buffer salts may begin to precipitate out of solution. For an ethanol-containing final solution, ethanol may be added up to 50% in the coupling buffer and still maintain buffer solubility. For ligands that are particularly insoluble in aqueous environments, there may be a micro-precipitate in the final solution, but the reaction still should proceed without difficulty.

3. Mix the washed gel with the ligand solution with stirring and add 0.3 g of EDC and mix to dissolve.



4. React with constant mixing for 2 to 4 h at room temperature. The gel slurry may be placed in a sealed container and mixed by end-over-end rocking.

5. Unreacted amines on the support may be blocked by the addition of a small hydrophilic carboxylate compound, such as glyceric acid (or 2,3-dihydroxypropanoic acid) or just by using sodium acetate. If a blocking step is desired, add the chosen blocking agent to the coupling reaction to make a final concentration of 0.1-M and readjust the pH, if necessary. Add an additional 0.3 g of EDC and react for 1 h with mixing.

6. Wash the support extensively with water, 1-M NaCl, and again with water to remove all unreacted components from the gel. Additional wash solutions may be utilized to completely remove ligands that may have some nonspecific binding potential to remain noncovalently bound to the immobilized ligand, such as the use of acidic and alkaline washes as well as washes containing denaturants (such as guanidine). After washing, the affinity gel may be stored as a 50% aqueous slurry containing a preservative at 4°C.

2.5. Streptavidin-Mediated Immobilization Methods

One of the most useful affinity interactions for bio-logical techniques involves the tight binding of biotin

(vitamin H) to the protein avidin (from egg whites) or streptavidin (from the bacterium Streptomyces avidinii). Both of these proteins are tetrameric in structure and each subunit contains a binding pocket for biotin. The deep binding pocket that biotin sits in contains amino acid residues able to form eight hydrogen bonding interactions plus additional van der Waals interactions with the bicyclic structure of biotin, thus creating one of the strongest noncovalent interactions known (dissocia-tion constant, (Kd): ~10−14–10−15 M). For this reason, the binding of biotin to avidin or streptavidin is similar to a chemoselective ligation reaction forming a covalent bond—highly specific and nearly irreversible.

Streptavidin conjugates are widely used as universal detection reagents in immunoassays, cellular imaging, flow cytometry, and other targeting and assay appli-cations. Conjugates of streptavidin with a fluorescent molecule or an enzyme can provide sensitive detection of biotinylated antibodies and other affinity targeting molecules (see Chapter 11). Immobilized streptavidin can also be used to capture biotinylated molecules out of complex solutions, providing a mechanism for isolat-ing interacting proteins or other biological complexes via immunoprecipitation (IP) (Figure 15.97).

For instance, an immobilized streptavidin support can be used to retrieve a biotinylated antibody that has specificity toward and has interacted with a desired protein target within a biological sample. As the affin-ity support captures the antibody–antigen complex

+

Biotinylatedantibody

SampleAntigen bound by

biotinylated antibodyin sample solution

Add immobilizedstreptavidin

S

S

Antigen isolatedon affinity support

FIGURE 15.97 Use of immobilized streptavidin for IP and co-IP applications.



through the biotin modifications on the antibody, it may also pull out of solution any proteins or other biomolecules that have interacted with the antigen (a technique sometimes called a “pull-down” assay). Co-immunoprecipitation (co-IP) often is used to study interacting proteins if the protein complexes have suf-ficient affinity to remain intact during the affinity sepa-ration process (see Chapter 24). After washing off any non-interacting proteins in the sample, the co-IP iso-lated complexes then can be eluted from the affinity support using a variety of conditions, which can con-sist of an acid pH buffer (e.g., 0.1-M glycine, pH 2.8) to break the antigen–antibody interactions or much stronger elution agents such as 8-M guanidine hydro-chloride, pH 1.5, or actually boiling the beads in SDS electrophoresis sample buffer, which will break the streptavidin–biotin interactions.

The preparation of an immobilized streptavidin affinity support can be achieved using amine-reactive resins, such as those described earlier in this chap-ter. For example, an aldehyde-containing matrix can be used to couple streptavidin by reductive amination and form a high-capacity affinity support for binding biotinylated proteins and other molecules. Coupling streptavidin to the support at a level of 5 to 10 mg/ml in the reaction medium will result in an excellent uni-versal affinity gel for the noncovalent immobilization of biotinylated antibodies. Many such streptavidin affin-ity supports are available commercially (e.g., Thermo Fisher, Sigma).

Immobilization of Biotinylated Antibodies on Immobilized Streptavidin

The following protocol is a generalized method for the immobilization of biotinylated antibodies on a streptavidin-agarose affinity gel. Some optimization of the loading level on the support may have to be per-formed to obtain the best performance of the coupled antibody in its intended application. Biotinylation of the desired antibody, if not commercially available, can be done using methods described elsewhere in this book (Chapter 11 and Chapter 18).

1. Wash 1.0 ml of an immobilized streptavidin support with water to remove storage solutions and then with 0.1-M sodium phosphate, 0.15-M NaCl, pH 7.2 (binding buffer). Washing may be carried out in a small column containing a bottom disk to prevent the gel from escaping. Wash with at least several column volumes. Drain to a wet cake and place the bottom cap on the column to prevent further flow.

2. Prepare a biotinylated antibody solution by dissolving it at a concentration of about 2 to 5 mg/ml in binding buffer. If the amount of antibody being immobilized is far less than this amount, then

it is best to use a proportionally lower amount of immobilized streptavidin.

3. Mix the biotinylated antibody solution with the washed streptavidin-agarose and stir to resuspend the gel in the solution. Place a top cap on the column and mix by rotation for 10 to 15 min at room temperature. For a reaction with microliter quantities of immobilized streptavidin, transfer the biotinylated antibody slurry to a small centrifuge tube, seal it, and mix by rotation for the recommended time.

4. Wash the immobilized antibody support extensively with binding buffer to remove any not-bound protein (at least 10 column volumes). After washing, the affinity gel may be stored as a 50% aqueous slurry containing a preservative at 4°C.

2.6. Protein A-, Protein G-, or Protein A/G-Mediated Antibody Immobilization Methods

The immunoglobulin binding proteins protein A, protein G, and protein A/G have been used extensively in immobilized form on chromatography supports for the purification of antibodies. Protein A is a 56-kDa cell wall constituent of the bacterium Staphylococcus aureus that contains five identical binding domains, each able to interact with the heavy chains of immunoglobu-lins in the region of the Fc fragment and in some cases within the region of the Fab fragments (Figure 15.98) (Graille et al., 2000; Idusogie et al., 2000). The recombi-nant form of protein A is typically truncated to about 45 kDa and is a robust, single polypeptide protein that can be immobilized onto chromatography supports through its lysine amine groups.

Protein G is another immunoglobulin binding pro-tein that originates in group C and G Streptococcal bac-teria. The native protein is expressed as a 56-kDa or 58-kDa polypeptide that contains multiple binding sites for immunoglobulins as well as a binding site for albu-min. The recombinant form of the protein is a truncated version that has the albumin binding site removed, but retains the IgG binding capabilities of the native mol-ecule. Protein G binds to antibodies through the heavy chains in the region of the Fc fragment, but at a differ-ent site than that of protein A. The differences in anti-body binding between protein A and protein G translate into differential binding specificities and affinities and thus offer options in binding and purifying antibodies, depending on the type of antibody desired.

A chimeric fusion protein consisting of the combi-nation of protein A and protein G, called protein A/G, merges the advantages of both protein specificities into one molecule. Protein A/G often is the immobilized immunoglobulin binding protein of choice for the puri-fication of a wide range of antibody types from various species.



However, the use of immobilized immunoglobu-lin binding proteins has been extended beyond just the affinity purification of antibodies. These supports can also be used to bind an antibody for subsequent targeting and immunoaffinity isolation of an anti-gen for which the antibody is designed to bind. Since most IgG type antibodies bind to protein A, G, or A/G in the region of their Fc fragments on the heavy chains, the antigen binding areas at the ends of the Fab frag-ments remain open to interact with antigens. This ori-ented binding of the antibody molecules places them in the ideal position to most effectively use both anti-gen binding sites and thus maximize the capacity of an immunoaffinity support. Other methods of antibody immobilization may result in almost random orien-tations of the antibody attached to the support, thus blocking some antigen binding sites by the antibody being positioned with the Fab ends facing the matrix instead of pointing out from it.

Schneider et al. (1983) described the use of immobi-lized protein A for the secondary immobilization of IgG type antibodies. Since the initial interaction of the antibody with protein A is potentially reversible just by changing the buffer conditions, this technique also uses a crosslinking agent to covalently trap the antibody on the support after binding to the protein A molecules. After applying the desired antibody to the protein A support and washing off excess not-bound material, the homobifunctional reagent DMP (dimethyl pimel-imidate; see Chapter 5, Section 2.2) is incubated with the bound antibody–protein A complex. The imidoester ends of this crosslinker effectively locks the antibody onto the support by reacting with lysine amines on both

the protein A and the antibody and forming covalent amidine bonds (Figure 15.99). This method has become quite popular but unfortunately it has one significant shortcoming: the amidine bonds formed from the reac-tion of the imidoesters with lysine amines are unstable and continually break down and leach antibody. This results in the presence of antibody in any isolated anti-gen preparations, which negatively affects the purity of most IP or co-IP experiments done.

Pierce (now Thermo Fisher) developed a modification of the Schneider method that solved the problem of con-tinually leaching antibody. Instead of using DMP with its imidoester reactive groups to lock in place the antibody–protein A interaction, the homobifunctional crosslinker DSS (disuccinimidyl suberate; see Chapter 5, Section 1.2) is used. The NHS ester ends of DSS are much more able to effectively crosslink and stabilize the protein A–antibody interactions, and since the resultant linkages involve the formation of amide bonds the stability of the immobilized antibody is excellent. Essentially no leach-ing of antibody molecules is observed using this tech-nique for IP or co-IP experiments. The crosslinker also stabilizes the antibody itself from breaking off the sup-port and releasing light or heavy chains during the elu-tion process from an experiment. In addition, this same crosslinking procedure can be used with immobilized protein A, protein G, or protein A/G supports to create immunoaffinity resins that are useful for the capture of any targeted protein or other antigen molecules from a complex biological solutions.

The downside of using an immunoglobulin-binding protein to immobilize an antibody with a crosslinker is that there may be nonspecific binding potentially

Protein Abindingdomain

Fabfragment

Protein Abindingdomain

Heavy-chain Fcregion

Carbohydratebetweenheavy chains

Antigenbindingregion

FIGURE 15.98 Three-dimensional molecular model of protein A binding to the Fc and Fab regions in antibodies (Graille et al., 2000; Idusogie et al., 2000; PDB IDs 1DEE and 1L6X.



created by the presence of the other components besides the antibody. Protein A, for instance, may have a tendency to interact with unwanted proteins in a sample, or the presence of crosslinker modifications may cause some additional nonspecificity especially if the cross-bridge is somewhat hydrophobic. In par-ticular, samples containing immunoglobulins should not be used with these supports, because the protein A on the matrix still has capacity to bind considerable IgG even with an antibody crosslinked onto its surface. Although the Schneider method and its various deri-vations are popular techniques to prepare an immuno-affinity support, if nonspecific binding is observed it may be advantageous to explore a method that allows direct coupling of the antibody to an amine-reactive, thiol-reactive, or carbonyl-reactive support as described previously in this chapter. Direct coupling would avoid any potential for nonspecific binding due to the protein A or the crosslinker used to stabilize the linkage.

Antibodies immobilized using a crosslinking process onto immunoglobulin binding supports have been used for many studies involving IP and co-IP procedures. Rubenstein et al. (2011) used an immobilized primary antibody on protein A/G–agarose to investigate the reg-ulation of endogenous ENaC functional expression by CFTR (cystic fibrosis transmembrane conductance reg-ulator) in airway epithelial cells. Qian et al. (2011) cou-pled a primary antibody to immobilized protein G on agarose to study the regulation of the alternative splic-ing of tau exon 10 by SC35 and Dyrk1A.

Immobilization of Antibodies on Protein A/G Supports

The following protocol is a generalized method for the binding and crosslinking of an IgG type antibody

(polyclonal or monoclonal) onto an immobilized pro-tein A/G support. Avoid the use of Fab fragments, as these will not interact as effectively with the protein A/G on the matrix surface, even though many Fab frag-ments do have a protein A binding site. Protein A/G is used instead of protein A due to its property of being a fusion protein and containing the combined properties of both protein A and protein G, thus providing excel-lent binding potential toward the widest variety of anti-body species and subclasses. This procedure is adjusted to be appropriate for the immobilization of 100 to 200 μg of IgG antibody onto 100 μl of immobilized protein A/G; however, the amount of resin prepared may be scaled up or down by proportionally changing the quantity of each reagent used. The protein A/G support material may be obtained commercially (e.g., Thermo Fisher) or it can be prepared by reacting 5 to 10 mg protein A/G per ml of an amine-reactive support. See the section in this chapter on amine-reactive immobilization methods for additional information.

1. Wash 100 μl of immobilized protein A/G with several volumes of water and then equilibrate the gel with coupling buffer (0.1-M sodium phosphate, 0.15-M NaCl, pH 7.2). The washing steps may be done in an appropriately sized drip column or spin column. If a spin column is used, avoid centrifuging at higher than about 100 to 300× g to prevent gel damage or clumping. Drain to a wet gel cake and place the bottom cap onto the column.

2. Prepare 100 μg of the antibody to be coupled at a concentration of 1 μg/μl in 100 μl of coupling buffer. If the antibody is already in solution in another buffer, dialyze or buffer exchange it into coupling buffer using size exclusion chromatography.

Immobilizedprotein A

IgG antibody

+

Affinity binding andcapture of antibody

on Fc fragment

Crosslinking ofantibody resultsin stable linkage

DSS

Orientedbinding

FIGURE 15.99 An immobilized protein A support can be first loaded with antibody through noncovalent affinity interactions with the Fc region of an IgG and then crosslinked with a homobifunctional reagent to stabilize the complex.



3. Mix the antibody solution with the immobilized protein A/G support with stirring to resuspend the gel. Place the top cap on the column or alternatively, transfer the slurry to a small tube and seal the tube with a cap. Mix the reaction medium with end-over-end rocking for 30 to 60 min to ensure complete binding of the antibody to the immobilized protein A/G.

4. Wash the support with 3 to 5 bed volumes of coupling buffer to remove any not-bound antibody and drain the gel to a wet cake.

5. Weigh 2 mg of the crosslinker DSS into a microfuge tube and dissolve it in 217 μl of dry DMSO or DMF to prepare a 25-mM solution. Make a 1 : 9 dilution with solvent by taking 20 μl of the DSS solution and diluting it with 180 μl of additional solvent to result in a 2.5-mM solution.

6. Add 300 μl of coupling buffer to the washed gel cake and mix to resuspend the gel. With mixing, add 100 μl of the diluted DSS solution to the gel slurry. The total slurry volume will be 500 μl, including the hydrated matrix. This mixture makes a final DSS concentration of 500 μM in the reaction, which will be sufficient to crosslink and stabilize the antibody onto the protein A/G attached to the resin.

7. React with constant mixing for 30 to 60 min at room temperature.

8. After the reaction is complete, wash the immunoaffinity resin with an elution buffer designed to remove antibodies from protein A/G affinity supports. A suggested buffer is 0.1-M glycine, pH 2.8. The acidic pH will break any remaining noncovalent protein A/G–antibody interactions and thus elute off non-crosslinked species. Wash with at least 5 column volumes of elution buffer to fully remove non-crosslinked antibody.

9. Wash the immunoaffinity support with water, 1-M NaCl, and water to completely remove the remaining buffers and reactants. The affinity gel may be stored as a 50% aqueous slurry containing a preservative at 4°C.

2.7. Reactive Hydrogen-Mediated Immobilization Methods

The methods commonly used to immobilize pro-teins or other biomolecules as well as couple organic compounds to chromatography supports usually relate to the covalent attachment of the ligand through a functional group, which most often includes amines, thiols, carboxylates, aldehydes, ketones, and occasion-ally hydrazides or aminooxy groups. However, some-times a ligand does not contain any of these standard functional groups to facilitate easy immobilization using the activation methods thus far described in this

chapter. In such cases, either the ligand must be deriva-tized to provide a functional group appropriate for coupling or another option must be used to facilitate immobilization.

One potential route to coupling ligands that do not have this common set of functionalities is to use a reac-tive hydrogen-mediated method, which targets certain replaceable hydrogen atoms within the compound’s structure. These methods may work for compounds containing reactive aromatic hydrogens that may be present within the ligand structure, as is often the case with certain drugs, steroidal compounds, dyes, or other aromatic organic compounds.

The following sections describe two reaction strate-gies that might be used to immobilize ligands having none of the standard functional groups, as described in the previous sections of this chapter.

Diazonium ActivationDiazonium reactions have been used for many

years in organic synthesis as well as for protein modi-fication, crosslinking, and immobilization procedures (Higgins and Fraser, 1952; Phillips et al., 1965; Inman and Dintzis, 1969; Cuatrecasas, 1970). Some of the earli-est dyes contained diazo groups within their structure, which were formed from the reaction of a diazonium intermediate with a reactive hydrogen on another aro-matic compound. The preparation of reactive chroma-tography supports containing diazonium groups can be done using an intermediary p-aminobenzylalkyl group, which in turn is created from the prior immobilization of a p-nitrophenyl derivative. A sequence of reactions is needed to form the required intermediate aminophenyl groups and then activate them to the diazonium deriva-tives. For immobilization reactions on chromatogra-phy supports, the diazonium groups need to be made immediately before use due to their high reactivity and instability.

Two routes to the formation of a reactive diazonium group on a support are shown in Figure 15.100. In one option, an amine-terminal spacer arm on a support is reacted with p-nitrobenzoyl chloride (Sigma-Aldrich) under nonaqueous conditions to create a nitrophenyl intermediate. In the second reaction option, the modi-fication reagent N-succinimidyl-p-nitrophenylacetate (SNPA; Bachem) is coupled to the amine-terminal spacer arms on the support to form an amide bond link-age, which terminates in the nitrophenyl groups. In both options, the nitro groups are then reduced to aro-matic amines through treatment with sodium dithionite in aqueous solution to form the necessary aminophe-nyl derivatives. This intermediate is stable for long-term storage until the support is needed to immobilize an affinity ligand. Just before the coupling reaction, the support is activated using an ice-cold solution of



sodium nitrite in HCl to create the reactive diazoniums from the aminophenyl groups. The reaction is done in the cold to prevent the diazonium group from immediately reacting with water to form phenol and nitrogen gas.

Diazonium groups are relatively unstable and should be used immediately to couple a ligand. The rate of diazo bond formation is extremely rapid with ligands, but even if no ligand is added to the support, diazo bond formation can be observed to occur within the matrix due to intramolecular crosslinks. This is due to the reaction of a diazonium group with the aminophe-nyl precursor molecules and will result in extensive

crosslinking and inactivation of the activated support within about an hour after activation. For this reason, ligand should be added quickly to limit the potential for the crosslinking side reactions.

Histidine and other imidazole-containing com-pounds can be coupled to a diazonium-activated sup-port at pH 8, while tyrosine and phenolic compounds are best immobilized in the range of pH 8 to 10. Figure 15.101 shows the reactions associated with ligand immobilization to create the final diazo linkage with the support. Ligands coupled using this method often result in a highly colored resin due to the presence of the diazo bonds. During the reaction, the support

NH2


++

N+

O–

O

Cl

O

p-Nitrobenzoylchloride

N+O–

O

O

ON

O

OSNPA

NH

ON+

O–

O N+O–

O

NH

O

Sodiumdithionite

Sodiumdithionite

NH

ONH2

NH2

NH

O

NaNO2,HCl

NaNO2,HCl

NH

ON N

Diazonium-activatedsupport

N

NH

ON


Cold Cold

FIGURE 15.100 Modification of an amine-containing support with p-nitrobenzoyl chloride or SNPA with subsequent reduction with sodium dithionite to form aryl amines. These intermediates can then be treated with sodium nitrite in cold, acidic conditions to create reactive diazonium groups.



can turn dark-brown or even black as the ligand mol-ecules are coupled. This color is normal and should not be viewed as unusual or a problem for subsequent use of the support for affinity separations. The diazo bonds also can be cleaved resulting in bond breakage and release of the immobilized ligand from the sup-port. Treatment with 0.1-M sodium dithionite in 0.2-M sodium borate, pH 9.0, results in complete bond cleav-age as evidenced by the disappearance of the diazo bond color within the support.

PREPARATION OF DIAZONIUM-ACTIVATED SUPPORTS

The following methods assume that an amine-con-taining support has previously been prepared accord-ing to the procedures outlined in the last section of this chapter. All operations should be done in a fume hood while wearing the appropriate personal protec-tive equipment to prevent contact with chemical com-pounds and solvents.

(A) Preparation of Aminophenyl Groups on the Support1. Wash 10 ml of an amine-containing resin with water

to remove any storage solutions and then wash the support into DMF (dry) using sequentially increasing concentrations of DMF in water (e.g., 30%, 60%) until 100% DMF is used. Washing can be performed using

a sintered glass filter funnel suspended in a vacuum filter flask in the fume hood. Continue to wash with at least 10 to 20 bed volumes of DMF until all remaining water is removed. Drain the support to a moist cake.

2. Prepare a solution of SNPA (Bachem) in DMF by dissolving 0.3 g of the modification reagent in 10 ml solvent. Mix thoroughly to dissolve.

3. With stirring, add the gel cake to the SNPA solution and mix the support to create a uniform suspension.

4. React with constant mixing for 1 h at room temperature. The mixing may be carried out in a sealed centrifuge tube by rotating it on an end-over-end mixer.

5. Wash the support extensively with DMF to remove excess reactants and reaction byproducts (at least 10 column volumes). Next, sequentially wash the support back into water by using decreasing concentration of DMF in water until pure water is used. Continue to wash with water (at least 10 column volumes) and then wash the support with several volumes of 0.1-M sodium borate, pH 9.0. Drain to a moist cake.

6. Prepare a solution of sodium dithionite in 0.1-M sodium borate, pH 9.0, by dissolving 1.2 g in 10 ml of buffer. Mix thoroughly to dissolve.

7. Add the moist gel cake to the dithionite solution with mixing. The dithionite will reduce the nitrophenyl

NH

ON N


+

+

HO

Tyrosine-or phenol-containing

ligand

NHN

Histidine-or imidazole-containing

ligand

NH

ON N

NH

ON N

HO

NHN

Immobilized tyrosinevia azo bond

Immobilized histidinevia azo bond

FIGURE 15.101 Immobilization of histidine (imidazole) or tyrosine (phenolic) ligands onto diazonium supports.



groups on the support to aminophenyl groups, the precursors used to form diazonium groups. React for 1 h at room temperature.

8. Wash the support thoroughly with 0.1-M sodium borate, pH 9.0, and then with water to remove all remaining reactants. The gel may be stored at this point before activation and coupling of ligand if desired. Storage should be done at 4°C as an aqueous slurry in the presence of a preservative.

(B) Activation to Form Diazonium Groups and Coupling of Ligands

Note that the following activation steps should be performed using ice-cold reagents and the reactions performed at 4°C. Do not allow the activated support to sit around for any length of time before coupling ligand, as the diazonium groups will quickly degrade and the support will lose coupling capacity. For this reason, the ligand solution (step 6) should be prepared prior to beginning the activation procedure.

1. Wash 10 ml of the aminophenyl-support prepared in (A) with ice-cold water and then with several bed volumes of cold 0.3-N HCl. Finally, suspend the gel in 10 ml of cold 0.3-N HCl using a 50-ml centrifuge tube or small vessel that can be sealed. Maintain the solution on ice.

2. Prepare 2.5 ml of a solution consisting of sodium nitrite (NaNO2) in cold water by dissolving it at a concentration of 50 mg/ml.

3. Add the sodium nitrite solution to the gel slurry with stirring.

4. React with mixing for 15 min at 4°C on ice.5. Quickly wash the activated gel with several bed

volumes of cold 0.3-N HCl, cold water, and then with cold coupling buffer (0.1-M sodium phosphate, pH 8.0, for coupling histidine/imidazole-containing ligands or 0.1-M sodium borate, pH 9–10, for coupling tyrosine/phenolic compounds). Drain the gel to a moist cake.

6. Prepare a ligand solution in the chosen coupling buffer (cold) at a concentration of 3 to 10 mg/ml for proteins or 3 to 5 mg/ml for small peptides or organic molecules. Some optimization of the ligand concentration should be carried out to obtain the best performance in the intended application. For ligands which are not very soluble in aqueous solution, the borate coupling buffer may be made with up to 50% ethanol (v/v) to aid in solubility. The ligand may first be dissolved in ethanol and then diluted in coupling buffer to promote solubility in the reaction medium.

7. Immediately add the ligand solution to the washed, activated support with stirring to resuspend the gel. React overnight at 4°C in a sealed container with constant mixing (e.g., end-over-end rocking).

8. Wash the affinity support with coupling buffer (containing ethanol if that was used to aid in solubility during the reaction) and then with water (again containing ethanol, if used), and finally with water alone. The affinity support can be stored until use at 4°C as a 50% aqueous slurry in the presence of a preservative.

Mannich CondensationThe methods available to immobilize molecules are

highly diverse and well characterized for ligands that contain a common functional group, which can easily be targeted and covalently linked to a support. Ligands that have at least one amine, carboxylate, aldehyde, ketone, thiol, or hydroxyl group can be coupled to a reactive solid support using one or more of the appro-priate reactions discussed previously in this chapter. However, for molecules that possess no such function-alities the route toward successful immobilization may not be as clear. In some cases, ligands such as drugs, steroidal compounds, inhibitors, dyes, or other organic molecules simply do not have a convenient functional group for linking to a support. In other cases, functional groups that are present may have low reactivity or be sterically hindered to allow efficient coupling to an acti-vated matrix.

Frequently, however, such difficult-to-immobilize molecules may contain reactive hydrogens (i.e., replace-able) within their structures to facilitate coupling in a Mannich condensation procedure. The classic Mannich reaction involves the condensation of formaldehyde (or another aldehyde compound) with ammonia (as its salt), and a third compound containing an active hydro-gen. The product of this reaction, usually performed under acidic conditions, involves the replacement of the active hydrogen with the methylene group from formaldehyde, and then attached to this is an amine, which originates from the ammonia. Instead of using ammonia, the reaction can also be performed using an amino group (primary or secondary) to achieve the linkage of the amine-containing compound with the active hydrogen-containing molecule through the meth-ylene (CH2) bridge. An example of this reaction using acetophenone, formaldehyde, and an organic amine salt proceeds as follows (the active hydrogens shown in bold text):

C H COC CH O RNH HCl C H COCH CH NH R HCl6 5 3 2 2 6 5 2 2H ⋅ → ⋅



Hermanson et al. (1992) first described the use of the Mannich reaction to immobilize ligands that did not have any common functional groups, but had suf-ficiently active hydrogens to participate in this reaction. Chapter 19, Section 6, also describes the use of this reac-tion for the bioconjugation of active hydrogen-contain-ing haptens to carrier proteins to produce immunogens suitable for immunization. The same reactions can be used to immobilize ligands onto amine-containing sup-ports using formaldehyde as the third reactant.

The use of the Mannich reaction to couple active hydrogen-containing ligands assumes the prior prep-aration of a support that contains spacer arms that terminate in the primary amino groups required to participate in the process (see the final section in this chapter for spacer arm preparation). The coupling reac-tion is driven in aqueous buffer by mildly acidic con-ditions and moderate heat, which most ligands of this type can tolerate without degradation. In addition, since one component of the reaction is already cou-pled to the support, no potential is present for uncon-trolled polymerization of the ligand in solution, as is often the case with molecules containing more than one reactive hydrogen undergoing Mannich condensa-tion. Figure 15.102 illustrates the immobilization of a steroidal molecule onto an amine-containing support

using this reaction process. The exact point of coupling to the ligand will be the most reactive or most replace-able hydrogen; however, there may be more than one orientation of ligand attachment if more than one active hydrogen site is present.

A modification of a Mannich coupling reaction for biomolecules was made by Joshi et al. (2004). This reac-tion uses aniline as the amine component similar to the methods used to enhance oxime or hydrazone for-mation (see previous sections, this chapter). However, instead of being a catalyst as it is in the reactions of aldehydes with hydrazides or aldehydes with amino-oxy compounds, in the Mannich reaction aniline actu-ally becomes incorporated into the final product. It first reacts with the formaldehyde (or another aldehyde) component to create an intermediate imine, which then goes on to specifically react with phenolic molecules in solution at positions ortho or para to the –OH group (Figure 15.103). The result is much more efficient end product formation even at room temperature than the standard Mannich reaction procedure, which is a ben-efit for coupling sensitive biomolecules.

This bioconjugation method of aniline-promoted Mannich condensation may also be extended to the immobilization of phenolic molecules by using immo-bilized aniline as the reactive group on the support. An

NH2


+

HO

OH

Estradiol-17beta

H2CO

H+, HeatNH

HO

HO

Immobilized estradiolvia a secondaryamine linkage

FIGURE 15.102 Immobilization of estradiol-17beta onto an amine-containing support using the Mannich reaction.

+NH2H2CO

NCH2

HO

NH

HO

Aniline Imineintermediate

Phenoliccompound

Conjugation atortho position

on phenol group

FIGURE 15.103 Mannich condensation reaction using aniline.



aniline-modified support was prepared in the previous section as a precursor to a diazonium-activated support. That same aniline intermediate may also be used in the Mannich reaction to couple with active hydrogens on phenolic compounds for the preparation of affinity chro-matography supports in a mild reaction process. Indeed, even proteins or peptides containing a tyrosine residue may be coupled specifically through the phenolic side chain using this reaction without affecting any other functional group on the molecule. The only requirement for proteins to efficiently participate in this reaction is that the tyrosine groups must be surface accessible. The reaction proceeds at room temperature and is optimal at a pH range of 5.5 to 6.5, which is mild for most biomol-ecules. Figure 15.104 illustrates the immobilization of a tyrosine-containing peptide on a support containing ani-line groups using the Mannich reaction.

Therefore, using the Mannich reaction two potential reaction paths are possible: (1) coupling of active-hydrogen containing molecules onto aliphatic amine-containing sup-ports when a ligand contains no other available common functional groups to facilitate coupling; and (2) immobi-lization of tyrosine-containing proteins or peptides onto aniline-containing supports specifically through tyrosine’s phenolic side chain while avoiding the reaction of other functional groups. These choices make the Mannich reac-tion a powerful alternative for the immobilization of cer-tain affinity ligands.

Because of the versatility of the Mannich conden-sation method of immobilization it is being used

more frequently for reactive hydrogen-containing ligands. Pyell and Stork (1992) used the method to prepare immobilized 8-hydroxyquinoline on an ami-nopropyl silane-modified silica support for use as a chelating agent. In addition, the phenol derivatives 4-(2-pyridylazo) resorcinol, 8-hydroxyquinoline, and 1-(2-pyridylazo)-2-naphthol were immobilized onto an amino-silica particle using the Mannich reaction (Tertykh et al., 2000), while Pu et al. (1998) used a simi-lar reaction process to couple 2-mercaptobenzothiazole to aminopropyltriethoxysilane-modified silica gel. In addition, Zhang et al. (2009) used the Mannich reaction to immobilize phenolphthalein onto an amine-modified polyacrylonitrile fiber for use as a halochromic fiber, which would change color with rapid response time depending on the pH of the environment. Members of this same group also used the Mannich reaction to produce a heavy-metal-detection fiber by coupling 4-(2-pyridylazo)-1,3-benzenediol onto ethylenediamine-modified polyacrylonitrile fibers (Li et al., 2010).

COUPLING LIGANDS VIA MANNICH CONDENSATION

The following protocols describe the use of either an aliphatic amine-containing matrix to immobilize active-hydrogen-containing ligands or an aniline-containing support to couple tyrosine-containing (or phenolic) ligands. Both reactions involve the use of formaldehyde and should be carried out in a fume hood using the appropriate personal protective equipment to prevent contact with reactants or solvents.

Aniline-containingsupport

+NH2H2CO

NCH2

Intermediateimine

NH

OHN

O

HO

Tyrosine-containingpeptide or protein

NH

NH

OHN

O

HO

Immobilization at tyrosineresidue via secondary amine linkage

FIGURE 15.104 Immobilization of a tyrosine-containing peptide on a support containing aniline groups using the Mannich reaction.



(A) Mannich Reaction with Active Hydrogen-Containing Ligands

This protocol assumes the prior preparation of an aliphatic amine-containing support as described in the last section of this chapter on the modification of supports with spacer arms.

1. Wash 10 ml of a primary amine-containing support with water to remove any storage solution and then with 0.1-M MES, pH 4.7 (coupling buffer). Examples of primary amine-containing spacer arm gels that work well in this process include MANAE–agarose and DADPA–agarose. Wash with at least 3 to 5 bed volumes of the solutions. The washing steps may be carried out in a small sintered glass filter funnel suspended in a suction filter flask. After washing, drain the support to a moist cake.

2. Dissolve the ligand containing an active hydrogen site in 10 ml of coupling buffer. If the ligand is relatively insoluble in aqueous buffer, it may first be dissolved in ethanol and then an aliquot added to the final buffer solution, making the total volume up to 10 ml; however, the final concentration of ethanol in the MES buffer should not exceed 50% to avoid buffer salt precipitation.

3. Mix the ligand solution with the washed amine-support and stir to resuspend the gel. Place the slurry in a sealable container and in a fume hood add to it 1.0 ml of 37% formaldehyde solution with mixing. Seal the container.

4. React at 37 to 57°C with constant mixing for a minimum of 24 h. For compounds with a slower reaction rate in the Mannich reaction, a longer reaction time or a higher temperature may be required for effective coupling yields. Some degree of optimization may have to be done to determine the best reaction conditions for a given molecule.

5. Wash the support extensively with coupling buffer and then with water (both containing ethanol, if used) to remove excess unreacted ligand. If the ligand is especially soluble in ethanol, a wash with 100% ethanol may be done after the water/ethanol wash to completely remove the last traces of free ligand. Finally, wash sequentially back into water and then with pure water. The affinity support may be stored as a 50% aqueous slurry at 4°C containing a preservative.

(B) Mannich Reaction with Tyrosine-Containing Ligands

This protocol assumes the prior preparation of an aniline-containing support as described in the previ-ous section on diazonium activation. Note that an alter-native approach to creating an immobilized aniline

intermediate is to immobilize 2-(4-aminophenyl)ethyl-amine directly onto an amine-reactive support through the aliphatic amino end. See the previous sections in this chapter on the preparation and use of amine-reactive supports for information on how to prepare this type of aniline-containing resin.

1. Wash 10 ml of an aniline-containing support with water to remove any storage solution and then with 0.1-M MES, pH 6.0 (coupling buffer). The washing steps may be done in a small sintered glass filter funnel suspended in a suction filter flask. After washing, drain the support to a moist cake.

2. Dissolve the ligand containing a tyrosine residue (i.e., protein, peptide, or a phenolic compound containing a free ortho or para position relative to its –OH group) in 10 ml of coupling buffer. If the ligand is relatively insoluble in aqueous buffer, it may first be dissolved in ethanol and then an aliquot added to the final buffer solution, making the total volume up to 10 ml; however, the final concentration of ethanol in the MES buffer should not exceed 50% to avoid buffer salt precipitation.

3. Mix the ligand solution with the washed aniline-support and stir to resuspend the gel. Place the slurry in a sealable container and in a fume hood add formaldehyde to make a final concentration of 25-mM. Mix well and seal the container.

4. React at room temperature (i.e., 20–25°C) with constant mixing for a minimum of 24 h.

5. Wash the support extensively with coupling buffer and then with water (both containing ethanol, if used during the coupling reaction) to remove excess unreacted ligand. If the ligand is especially soluble in ethanol, a wash with 100% ethanol may be done after the water/ethanol wash to completely remove the last traces of free ligand. Finally, wash sequentially back into water and then with pure water. The affinity support may be stored as a 50% aqueous slurry at 4°C containing a preservative.

2.8. Creating Spacer Arms on Chromatography Supports

The most common route of preparing an affinity chromatography support involves the direct activation of a solid-phase material followed by the coupling of the affinity ligand. Most of the methods described in the previous sections of this chapter use this path to ligand immobilization. Occasionally, however, it is useful to create an intermediate derivative on the support that includes the formation of a spacer arm, which might terminate in a functional group useful for subsequent reactions. Spacers are typically low-molecular-weight molecules that are most often linear and consist of



alkyl or hetero-alkyl components that extend out from the matrix from about 2 to 20 atoms in length. Spacer arms can serve multiple purposes on chromatography supports. They can simply be a bridge between an ini-tial functional group or reactive group on a support and the creation of another needed functionality at the other end of the spacer. They also can provide a long tether that extends the affinity ligand out from the matrix sur-face, thus providing greater steric accommodation for target molecules that may interact or dock with it. This is an especially important feature if the binding site for a ligand is a deep pocket within the target protein, such as is the case for avidin or streptavidin in binding to immobilized biotin. Even with much larger ligands, however, spacers can be used to extend the molecule away from the matrix surface for better accessibility to bind targets. Extremely long spacers such as PEG-based compounds can provide much greater freedom of motion for immobilized proteins, thus allowing greater potential for interactions to occur within the support. This may be especially important if an immobilized protein has more than one binding site for a target mol-ecule, as is the case for many immunoglobulin-binding proteins (i.e., protein A, protein G, and protein A/G) or in the case of avidin or streptavidin used to capture bio-tinylated proteins.

Spacers can also create additional properties on a support, such as the introduction of increased hydro-philicity that can reduce nonspecific binding to the final affinity resin. An early report recognized the benefit of using hydrophilic linker arms in the design of affinity chromatography supports (O’Carra et al., 1974). Spacers can also consist of branched constructs that create multi-point immobilization sites for an affinity ligand, which can stabilize some proteins or enzymes (e.g., gly-cidol modified supports, see previous corresponding section, this chapter). If desired, the cross-bridge of a spacer arm can also be chosen to be cleavable to permit chemical release of an affinity ligand after it has bound a target. Sometimes, in the design of an affinity support, it takes one or two spacer arm modifications to create the desired effects and functionalities on the matrix prior to the immobilization of the affinity ligand. In other cases, a spacer is formed during matrix activation or coupling of a ligand, such as in activation methods using a bis-epoxide compound or in the immobilization of an amine-containing ligand to an azlactone-activated support (see previous sections, this chapter).

The options available for adding spacer groups onto chromatography supports are numerous and mixing and matching these options can multiply the design choices dramatically. In the most fundamental spacer arm design, there are functional groups on both ends of a molecule that are separated by an aliphatic or hetero-aliphatic chain. Less frequently, there might be aromatic

groups present in the bridging portion of the spacer, but this is less common due to the hydrophobicity such groups can add to the final support. Spacers can even be constructed from the use of homobifunctional or het-erobifunctional crosslinking agents through the reac-tion of one end of the crosslinker to a functional group on the surface of the support, which leaves the other reactive group free to couple with a ligand or another spacer molecule. The epoxy activation of a support using the bis-epoxide compound 1,4-butanediol digly-cidyl ether yields a hydrophilic 12-atom spacer, which terminates in a reactive epoxide group before a ligand is coupled.

It is also important to consider the overall chroma-tography effects in having a spacer arm in the design of an affinity resin. Some of the potential consequences of an inappropriate linker arm include the introduction of hydrophobic or ion-exchange interactions that can result in binding to nonrelevant molecules in a sample, resulting in lower purity for the desired isolated target molecule. For instance, long aliphatic chains can cre-ate considerable hydrophobicity on an otherwise fairly hydrophilic base support. Although spacer arms of long length may seem desirable, increasing the length of an aliphatic spacer may only serve to increase the hydro-phobic interaction potential of the final affinity resin. A better alternative would be to choose a hydrophilic spacer instead of a hydrophobic one, which may pro-vide the extended length needed while still maintaining or even improving the hydrophilicity, and therefore low nonspecific binding character, of a support. Similarly, some spacers may create charges on a support from ion-ized or protonated groups, thus generating the poten-tial for nonspecific binding with oppositely charged molecules in a sample. Secondary amine-containing spacers are particularly notorious for creating posi-tive charge on a matrix, which may lead to undesirable interactions. Spacers containing carboxylates or sulfo-nates may also generate nonspecific binding character-istics from creating negative charges on the support.

Figure 15.105 shows examples of common spacer molecules used in the preparation of immobilized affin-ity ligands, which is by no means exhaustive. Many of these molecules contain aliphatic or hydrophobic cross-bridges, which should be used with caution to avoid nonspecific binding issues in the final support. Even the spacers containing only a six-carbon aliphatic bridge can result in considerable hydrophobic interac-tion potential, even though they are used quite fre-quently with success in designing affinity supports. For hydrophilicity in the final support, it is impor-tant to choose a spacer composition containing polar groups within an alkyl chain, which can reduce or eliminate hydrophobic character. The best choices for water solubility and low nonspecific binding include



polar components such as secondary amines, carbonyl groups, amide bonds, ether groups, and hydroxyls. If possible, avoid charged groups, such as protonated secondary (or tertiary) amines or carboxylates, which may introduce ion-exchange effects. Perhaps a supe-rior choice in this regard is to use a PEG-based spacer,

which avoids both the issue of hydrophobicity and also does not generate within the cross-bridge any charged sites with ion-exchange potential. There are now many discrete PEG-based spacers and crosslinkers available for the design of virtually any immobilized ligand (see Chapter 18). A list of spacers and modification reagents

H2NNH2

Ethylenediamine

NH2H2NOH

1,3-Diamino-2-propanol

HN NH2H2N

Diaminodipropylamine

H2NS

SNH2

Cystamine

NH2H2N

1,6-Diaminohexane

H2NO

ONH2

Jeffamine EDR-148

H2NO

OO

NH2

Jeffamine ED-600

x y z

OO

OH2 NN H2

4,7,10-Trioxa-1,13-tridecanediamine

H2NO

OO

OO

OO

OO

OO

NH

O

O

Boc-N-amido-dPEG11-amine

H2N

O

OH

beta-Alanine

NH2HO

O

Aminocaproic acid

H2NO OH

O

nAmino–PEGn–carboxylate

O

HOO

OH

O

OH

O

HOO

O

OHDiglycolic acid

HO

OHS

O

OHThioglycolic acid

N

O

O

O

OS

OSATA

N

O

OO S

O O

SATPO

OOO

ON

O

O

OS CH3

O

SAT–PEG 4

NH

OO S

N-Acetyl homocysteinethiolactone

O

OHHS

8-Mercaptooctanoic acidSS

O

OH

Alpha-lipoic acid

SS

O

HN

O OHn

Lipoamide–PEGn–carboxylate

HSOH

OHSH

DTT

Tetra(ethylene glycol) dithiol

HSO

OO

OO

OSH

Hexa(ethylene glycol) dithiol

H2NSH

2-Mercaptoethylamine

O

OHN

H2N NH

NH2

Adipic dihydrazide

HN

NH2

O

HN

H2N

Carbohydrazide

Succinic acid

Glutaric acid

O

HSO

OO

OSH

FIGURE 15.105 Small molecules used as spacers in affinity chromatography.



that have been used to create linker molecules on solid phases and categorized by the type of spacer follows.

Diamine SpacersEthylenediamine1,3-Diamino-2-propanolDADPACystamine1,6-DiaminohexaneJeffamine EDR-148Jeffamine ED-6004,7,10-Trioxa-1,13-tridecanediamineBoc-N-amido-dPEG11-amineBoc-N-amido-dPEG3-amine

Amino–Carboxylate SpacersBeta-alanineAminocaproic acidAmino–PEGn–carboxylate compounds

bis-Carboxylate SpacersSuccinic acid (or succinic anhydride)Glutaric acid (or glutaric anhydride)Diglycolic acid (or diglycolic anhydride)

Thiol–Carboxylate SpacersThioglycolic acidSATASATPN-Acetyl homocysteine thiolactone8-Mercaptooctanoic acidAlpha-lipoic acidLipoamide–PEGn–carboxylate compoundsThiol–PEGn–carboxylate compoundsNHS–PEGn–acetylated thiol compounds (SATA-like)

bis-Thiol SpacersDTTTetra(ethylene glycol) dithiolHexa(ethylene glycol) dithiolPoly(ethylene glycol) dithiol

Amino–Thiol Spacers2-mercaptoethylamine

bis-Hydrazide SpacersAdipic dihydrazideCarbohydrazide

Of course, some hydrophobic spacer molecules may be purposely chosen to result in a resin having hydro-phobic character designed into the gel for certain sep-arations. Various forms and lengths of hydrophobic linkers can be coupled to a support to create a resin useful for hydrophobic interaction chromatography. In this case, the longer the hydrophobic spacer arm, the greater will be the hydrophobic interaction potential. A 4-carbon chain, for instance, will have much weaker hydrophobic interaction potential with proteins than an 8-carbon chain. The extreme hydrophobicity of an 18-carbon aliphatic chain is often designed into HPLC

chromatography resins, which is the length typically used for reverse phase separations.

The following sections describe some of the most popular spacer molecules used to make affinity res-ins and the reactions that can be used to prepare the derivatized supports. The choice of spacer for a par-ticular immobilization reaction should be done based upon the functional groups required at both ends of the spacer and the length and physical properties of its cross-bridge. To design an optimal affinity resin containing a spacer molecule, it is often best to evalu-ate a number of spacer arm types to determine which one performs best for a given application. For instance, if there is a need for a diamine spacer in the construc-tion of an immobilized ligand, then it is best if several different diamines are compared to see how changes in spacer structure and length affect the final affinity separation.

Diamine SpacersSpacer molecules containing an amine at both ends

often are used to create a primary amine on a support for further modification or for the coupling of carbox-ylate-containing ligands. They are also used as an inter-mediary in the creation of some activated supports, such as in the preparation of an iodoacetyl-support useful for coupling thiol-containing ligands (see earlier section, this chapter). To couple a diamine spacer to a support, an amine-reactive resin should first be created according to the methods described previously. The diamine is then reacted in large excess with the amine-reactive support to result in one end of the spacer cou-pling to the support and the other end remaining free. To reduce the potential for crosslinking that results in both ends of the spacer being linked to the support, the reaction needs to be performed at a minimum concen-tration of 0.5-M diamine. Alternatively, if the diamine spacer molecule is available with one end protected (one amine blocked), then a much lower concentra-tion can be used for coupling; however, the final prod-uct then needs to be deblocked to reveal the terminal amine for further modification. Typical protecting groups for amines include carbobenzyloxy (Cbz), tert-butyloxycarbonyl (BOC), and 9-fluorenylmethyloxy-carbonyl (FMOC), which allow one end of the diamine to be immobilized without the potential for crosslink-ing. Removal of the protecting groups is by catalytic hydrogenolysis (Cbz), strong acid such as trifluoroace-tic acid (TFA) (BOC), or an organic base such as piperi-dine (FMOC). Use care to ensure that the deprotection step does not cause adverse effects on the base support. In this regard, deprotection of Cbz groups may not be compatible with porous chromatography supports, as the typical reagent used for this procedure is a particu-late Pd/C suspension and hydrogen. The Pd/C particles



may get entrapped within the matrix and not be easily removed. A better protecting group would be FMOC, because the addition of a 20% piperidine solution in DMF for the deprotection step would be tolerable for most support materials.

Diamine compounds that are in the free base form (not as salts) are extremely caustic in aqueous solution and upon dissolving will typically give a pH > 11. The dissolving process in water and the subsequent neu-tralization reaction with acid can be hazardous, as both are exothermic processes. If the dihydrochloride salt of a diamine is commercially available, then the neutral-ization step is not needed and the diamine typically is supplied as a solid and is much more convenient to work with. To neutralize diamines safely, always use a fume hood and proper personal protective equip-ment to avoid contact with solutions and inhalation of fumes. To limit the exothermic heat effect, a diamine compound may be first added to an excess of crushed, deionized ice (equal to a little less than the final desired solution volume) and then 6-N HCl added slowly with stirring. As the diamine is neutralized, the ice will melt and prevent the solution from becoming extremely hot. Once the diamine solution is neutralized, the appropri-ate buffer salt may be added and the pH adjusted to the proper value for the coupling reaction being done.

DIAMINODIPROPYLAMINE

One of the more common diamine spacer molecules used in affinity chromatography is diaminodipropyl-amine (DADPA), also known as 3,3′-iminobispropyl-amine (Figure 15.106). It is a nine-atom spacer with a central secondary amine and a primary amine at each of the ends. The central secondary amine aids in overall hydrophilicity, but may also be a potential site of posi-tive charge due to protonation at physiological pH. In addition, the secondary amine might participate in some reactions that are targeted at the terminal primary amine, thus complicating the structural nature of the final immobilized ligand. DADPA is liquid at room tem-perature (d = 0.938 g/ml; mp = −14°C) and volatile with a strong amine odor; therefore, all solutions should be handled in a fume hood. The reagent is soluble in aque-ous solution for buffered reactions and it is also soluble in many organic solvents suitable for work with chro-matography supports (i.e., DMSO, DMAC, DMF, ace-tone, dioxane, and ethanol).

Amine-reactive supports may be modified with DADPA to produce spacer arms terminating in a pri-mary amine. When using amine-reactive supports that are labile in aqueous environments (e.g., due to hydrolysis), then the DADPA coupling reaction can be done in organic solvent to eliminate the competing


O

H

O

CDI-activated support

O

N

N

H2N NH

NH2

H2N NH

NH2

+

+

HN

HN NH2

O

O

NH

NH

NH2

DADPA

DADPA

NaCNBH3

Coupling via secondaryamine linkage

Coupling via amide linkage

FIGURE 15.106 Coupling the diamine spacer DADPA to periodate-oxidized agarose and CDI-activated agarose.



hydrolysis reaction and maximize yields. DADPA reac-tions should be done with a large excess of diamine (at least 0.5-M) to limit the potential for crosslinking within the matrix and maximize the probability of only one end of each molecule reacting with the support.

Immobilized DADPA has been used to couple car-boxylate-containing antigens using the carbodiimide EDC, which resulted in an affinity support for the puri-fication of specific antibodies from immune serum (Tsai et al., 1998). It has also been used as a precursor in the preparation of a maleimide-activated support by react-ing the heterobifunctional crosslinker SMCC (Chapter 6, Section 1.3) with the amine on the support to yield a thiol-reactive derivative for the coupling of the rat GIP receptor epitope using a cysteine-terminal peptide. This affinity support was subsequently used to purify anti-bodies against the intact receptor protein (Lewis et al., 2000). Shenouda et al. (2002) similarly used immobilized DADPA to couple a hapten peptide sequence (human urotensin-II) to the support using the EDC amide bond forming method, and the affinity support then was used to purify anti-urotensin-II antibodies from rabbit antiserum. In addition, Kost et al. (2011) used DADPA–agarose to couple the CNS-modulating compound clavulanic acid via its carboxylate using EDC and subse-quently was able to isolate two proteins that specifically interacted with this ligand from neuronal cells.

PREPARATION OF A DADPA-MODIFIED SUPPORT The following protocols represent two methods of cou-pling DADPA to amine-reactive supports. Analogous methods may be used with other short diamine spac-ers, such as ethylene diamine or 1,3-diamino-2-propa-nol. In the first method, a periodate-oxidized agarose support is used in a reductive amination procedure in aqueous solution (see previous section on immobiliza-tion by reductive amination, this chapter). In the second protocol, DADPA is coupled to a CDI-activated support in a nonaqueous solution to eliminate the hydrolysis reaction, which would occur with the reactive support in aqueous solution. Both of these methods yield sup-ports containing spacer arms that terminate in free pri-mary amines for further reactions. The CDI coupling procedure will give a higher density of amines on the support, but both work quite well for subsequent immobilization reactions. All operations should be car-ried out in a fume hood.

(A) Coupling DADPA to Periodate-Oxidized Agarose1. Wash 100 ml of periodate-oxidized agarose (or

another aldehyde-containing support) with water to remove storage solutions and then into coupling buffer (0.1-M sodium phosphate, pH 7.2). Washing steps may be performed with a sintered glass filter

funnel suspended in a vacuum filter flask. Wash with at least several bed volumes for each wash step. Drain to a moist cake.

2. In the fume hood, add 20 g (21.3 ml) of DADPA to about 100 ml of crushed, deionized ice (use personal protective equipment to prevent contact with the highly caustic diamine and the acid used for neutralization). Slowly add 8 to 10 ml of concentrated HCl to the DADPA/ice slurry with manual stirring. Add the acid slowly using a pipette. As the acid is added, the solution will warm and the ice will melt. After the acid has been added, stir the solution using a magnetic stir bar and continue to neutralize the solution to about pH 7 with acid. Add to this solution a quantity of sodium phosphate buffer salt to make the final concentration 0.1-M phosphate when the total volume is diluted to 100 ml. Stir and readjust the pH to 7.2 using acid or base. The final solution is 1.52-M DADPA, 0.1-M sodium phosphate, pH 7.2. Allow the solution to come to room temperature before continuing.

3. Add the washed periodate-oxidized agarose to the DADPA solution and mix to resuspend the gel.

4. In the fume hood weigh out and add 0.63 g of sodium cyanoborohydride (toxic!) to the gel slurry and mix for 2 to 4 h. Mixing may be done using an overhead paddle stirrer (not a stir bar) or in a sealed plastic container by end-over-end rocking. Avoid the use of sealed glass containers, because there is some gas evolution during the reductive amination process.

5. Extensively wash the DADPA–agarose support with water, 1-M NaCl, and water to completely remove unreacted diamine and reaction byproducts. The support may be stored as a 50% slurry containing a preservative at 4°C until use.

(B) Coupling DADPA to CDI-Activated Supports1. In a fume hood, dissolve 20 g of DADPA (21.3 ml) in

100 ml of dry acetone with stirring (makes a 1.52-M DADPA solution).

2. Drain 100 ml of a CDI-activated support prepared in acetone of excess solvent using filtration on a sintered glass filter.

3. Add the drained support as an acetone-wet cake to the DADPA solution with mixing to resuspend the gel.

4. Stir the reaction slurry for 2 to 3 h using an overhead paddle stirrer or in a sealed container by end-over-end rocking.

5. Wash the modified support with 1 l of acetone to completely remove excess diamine. Next, sequentially wash the support into water by using increasing concentrations of water in acetone (e.g., 30%, 60%, and 100%). Continue to wash with water to completely remove the last traces of acetone, and



then with 1-M NaCl, and again with water. The support may be stored as a 50% slurry containing a preservative at 4°C until use.

1,6-DIAMINOHEXANE

The aliphatic diamine spacer 1,6-diaminohexane (DAH; also called hexamethylenediamine) has been used in a variety of applications in organic synthesis, including the formation of spacer arms on chromatog-raphy supports. DAH consists of a linear six-carbon linker arm with a primary amine on each end, and it is a solid at room temperature (mp 42–45°C) with hygro-scopic properties. The crystals of DAH should be white or off-white in color, but occasionally may have a yel-low tint to them. These are typically all right to use without further purification; however, avoid highly col-ored material as they are likely to be oxidized or con-taminated. Nearly one billion kilograms of DAH are produced annually as a key ingredient for several poly-mers. The use of DAH as a spacer arm for the prepara-tion of affinity supports dates to the earliest research on matrix preparation (O’Carra and Barry, 1972).

Since DAH does not contain any other polar con-stituents in its cross-bridge, it is considerably more hydrophobic than DADPA, discussed previously. For this reason, spacer arms built from DAH on chromatog-raphy supports will create more hydrophobic character on the matrix surface, even after an affinity ligand is attached. This can have detrimental effects on the spe-cific binding potential of an affinity support, because hydrophobic molecules in samples may bind through hydrophobic interactions and result in lower selectiv-ity for capturing the desired target molecule. In some cases, however, DAH has been found to be optimal in creating affinity supports with certain ligands immo-bilized, such as the dye Cibacron Blue 3GA (Suen and Tsai, 2000). In addition, for some applications of immo-bilized metal affinity chromatography (IMAC) it was found that medium-length hydrophobic spacers created a support with the best binding characteristics toward penicillin G acylase (Liu et al., 2005). Recent novel appli-cations of immobilized ligands in the field of supported photosensitizers (SPS) indicate that even longer ali-phatic spacer arms showed better efficiency than those with shorter chains (Pineiro et al., 2010).

DAH has also been used as a type of affinity ligand in the isolation of human IgG from serum and plasma, which involved a one-step purification process using HEPES buffer at pH 6.8 (de Souza et al., 2010). Bayramoglu et al. (2006) used DAH as a spacer on poly-mer beads containing epoxy groups to subsequently couple L-histidine as an affinity ligand for the purifica-tion of immunoglobulins.

DAH dissolved in aqueous solution is extremely caustic and should be handled with care. Upon

dissolution, the pH should be adjusted to the recom-mended point for coupling the spacer to an amine-reactive support and maintained with the appropriate buffer. Neutralization should be carried out using HCl and cooled on ice, similar to the procedure outlined in the previous section on DADPA. Alternatively, the dihydrochloride form of DAH can be used, which does not affect the pH of an aqueous solution, nor does it require titration with acid for neutralization.

PREPARATION OF A DAH-MODIFIED SUPPORT The following protocol describes the coupling of DAH to a tresyl- or tosyl-activated support, prepared according to the procedures described previously in this chapter (Figure 15.107). Other amine-reactive supports may also be used, such as the activated supports and general pro-cedures outlined in the section on the spacer DADPA.

1. In a fume hood, dissolve 15.28 g of DAH dihydrochloride in 100 ml of coupling buffer (0.2-M sodium phosphate, pH 7.5) and mix to dissolve. Adjust the pH back to 7.5 if necessary. Note: Alternatively, dissolve 10 g of DAH (as the free base) in a minimum quantity (~70 ml) of coupling buffer. Cool the solution in an ice bath. Slowly titrate the pH back down to 7.5 using concentrated HCl (caution: highly so use personal protective equipment!), while maintaining the solution cool to control the heat of the exothermic process. After the proper pH is reached, adjust the total solution volume to 100 ml by the addition of coupling buffer.

2. Drain 100 ml of a tresyl- or tosyl-activated support of acetone using a sintered glass filter suspended in a suction filter flask in a fume hood. Pull a gentle vacuum on the support and break up the matrix into small pieces using a spatula. Do not allow the support to dry. Remove the vacuum and add an equivalent bed volume of water and mix to resuspend the support. Continue to quickly wash the support with 3 bed volumes of water and 2 volumes of coupling buffer. Drain to a moist cake.

3. Mix the washed support with the DAH solution and mix to resuspend the gel.

4. If a tresyl-activated support was used to couple DAH, react with constant mixing for 2 to 4 h at room temperature. If a tosyl-activated support was used, the reaction should be continued overnight at room temperature to realize maximal coupling yields. Mixing may be performed using an overhead paddle stirrer or in a sealed container by end-over-end rocking.

5. Wash the support with coupling buffer, 1-M NaCl, and again with water to remove excess uncoupled DAH. The support may be stored as a 50% slurry containing a preservative at 4°C until use.



DIAMINE–PEG COMPOUNDS

Diamine spacers that contain a central PEG chain with amines on both ends are among the most hydro-philic bridges that can be created on a chromatogra-phy support. There are compounds available in this design that have a broad range of spacer arm lengths, from short PEG chains to those with high-molecular-weight long polymer chains. The discrete, short PEG compounds are useful for building amine functional-ity on a support while maintaining a hydrophilic and low nonspecific binding character on the base matrix. Longer PEG spacers might be chosen to further mask a more hydrophobic base support by developing a poly-ether coating that terminates in an amine. For the devel-opment of affinity supports for use with biomolecules, the choice of a diamine–PEG spacer may be an excellent starting point for high purity target molecule purifica-tion, as the spacer itself will not create hydrophobic or ionic interaction sites on the final support.

Diamine compounds containing a linear PEG bridge can be obtained from a number of sources. The short-est diamine–PEG spacers can be obtained from Aldrich [2,2′-(ethylenedioxy)bis(ethylamine)] or Huntsman (Jeffamine EDR-148) in the form of a PEG2 length and having ethylamine groups on each end. This compound is available in bulk quantities from Huntsman (from 5-gallon pails to tank-car loads), and Aldrich offers it in 100-ml and 500-ml package sizes, which are more

appropriate for small-scale affinity support production. This compound can easily be coupled to an amine-reac-tive support to yield a 10-atom spacer, which terminates in an amine. Another, slightly longer diamine–PEG compound is available from Aldrich that contains a PEG3 bridge with propylamine groups on each end (4,7,10-Trioxa-1,13-tridecanediamine). This compound is still extremely hydrophilic and provides a 15-atom spacer once it is coupled with an amine-reactive support (Figure 15.108). Both the PEG2 and PEG3 diamine spacers need to be reacted in high molar excess to avoid the potential for both ends of the molecules reacting with the activated support and crosslinking within the matrix.

Longer PEG-based diamine spacers can be obtained from Huntsman with the additional incorporation of a propylene oxide-capped polyethylene glycol internal construct. For instance, the Jeffamine ED series (e.g., ED-600) (Figure 15.109) contains a core PEG9 repeat with several propylene oxide groups at each end, which are terminally capped with a primary amine. The reagents are extremely hydrophilic and provide lon-ger PEG-based spacers at relatively inexpensive price points. These are good choices if a longer spacer arm is needed to move an affinity ligand away from the base matrix to provide better accessibility for the docking of biomolecules.

Discrete diamine PEG–compounds are also avail-able from Quanta Biodesign. Two of these spacers have

Tresyl-activated support


H2NNH2

+

NH

NH2

Coupling viasulfonamide linkage

Coupling viasecondary amine

linkage

OS

O

OCF3

DAH

H2NNH2

DAH

+

O SO

OCH3

OS

O

O HN NH2

FIGURE 15.107 Coupling the diamine spacer DAH to tresyl- and tosyl-activated supports.



one of their two amine groups protected to facilitate coupling to a support without the need to use a large molar excess to avoid crosslinking. Since only one amine will initially couple to the amine-reactive sup-port the final density of amines on the matrix can be finely controlled to an optimal level for immobilizing whatever affinity ligand is desired. This avoids hav-ing a high density of amines on the support that could contribute to an ion-exchange character, especially if the excess amines are not blocked after coupling a ligand. The particularly long protected diamine–PEG reagent t-boc-N-amido–dPEG11–amine contains a 37-atom cross-bridge that is extremely hydrophilic with a free amine on one end and a BOC-protected amine on the other end. The use of a protected diamine–PEG spacer does require a deprotection step after coupling to a support to free the protected amine for further modifications. The type of protecting agent used with these reagents

is a tert-butyloxycarbonyl (BOC) group, which typically requires strongly acidic conditions (TFA; 30–90%) in a water-immiscible solvent (dichloromethane) for depro-tection. Some supports may not be able to tolerate these conditions without some degradation and washing in and out of a water immiscible solvent is problematic. Some modifications to this standard scheme may work for aqueous phase deprotection. Heating to 60 to 100°C in a neutral pH water solution was found to be effi-cient in one study and may be viable for use with cross-linked chromatography supports (Wang et al., 2009). In addition, a modified acidic deprotection method was investigated by Han et al. (2001) that demonstrated fast deprotection using 4-N HCl in anhydrous dioxane. This method uses acidic cleavage of the protecting group, but it does so in a water-miscible solvent, which makes operations using chromatography supports more amenable to washing in and out of an aqueous envi-ronment. Two examples of discrete diamine–PEG com-pounds are shown in Figure 15.110, both of which use the BOC protecting group. The coupling and deprotec-tion reaction for the shorter compound is illustrated in Figure 15.111.

COUPLING A DIAMINE–PEG-BASED SPACER The following protocols describe the use of diamine–PEG-based spacers to modify amine-reactive sup-ports for additional reactions or for the coupling of


O

H

+

+

NaCNBH3


Coupling via amide linkage

H2N OO

O NH2

4,7,10-Trioxa-1,13-tridecanediamine

HN O

OO NH2

Suppport containingazlactone groups

HN

O

N

OO

O

H2NO

ONH2

Jeffamine EDR-148

NH

OO

NH2

FIGURE 15.108 Coupling of Jeffamine EDR-148 or the diamine–PEG3 compound to amine-reactive supports to yield terminal amine groups.

H2NO

OO

NH2

Jeffamine ED-600

x y z

y = 9 x + z = 3.6

FIGURE 15.109 Jeffamine ED-600 structure.



a carboxylate-containing ligand. The first procedure makes use of 4,7,10-Trioxa-1,13-tridecanediamine as the free amine (not protected and not as the dihydrochlo-ride salt) in the modification of an aldehyde-containing support using reductive amination (see discussion on reductive amination, this chapter). The second protocol makes use of a discrete diamine–PEG compound that has one end protected with a BOC group.

(A) Coupling of 4,7,10-Trioxa-1,13-tridecane-diamine to Periodate-Oxidized Agarose1. Wash 100 ml of periodate-oxidized agarose (or

another aldehyde-containing support) with water to remove storage solutions and then into coupling

buffer (0.1-M sodium phosphate, pH 7.2). Washing steps may be performed with a sintered glass filter funnel suspended in a vacuum filter flask. Wash with at least several bed volumes for each wash step. Drain to a moist cake.

2. In the fume hood, add 33.5 g (33.4 ml) of 4,7,10-trioxa-1,13-tridecanediamine (Aldrich) to about 100 ml of crushed, deionized ice (use personal protective equipment to prevent contact with the highly caustic diamine and the acid used for neutralization). Slowly add 8 to 10 ml of concentrated HCl to the diamine/ice slurry with manual stirring. Add the acid slowly using a pipette. As the acid is added, the solution will warm and the ice will melt. After the acid has

BoC-N-Amido–dPEG3–amine

H2N OO

OHN O

O

H2NO

OO

OO

OO

OO

OO

NH

O

O


FIGURE 15.110 Diamine–PEGn compounds with one end blocked by a BOC group.

Support containingNHS ester-reactive groups

O

ON

O

O

+


O

HN

Immobilization throughamide bond formation

H2N OO

OHN O

O

OO

OHN O

O

O

HN O

OO NH2

4 M HClin dioxane

Deprotection to primary amine

FIGURE 15.111 Coupling of a BOC-protected diamine–PEG compound to an amine-reactive support with subsequent deprotection with 4-M HCl in dioxane or heating to 60 to 100°C.



been added, stir the solution using a magnetic stir bar and continue to neutralize the solution to about pH 7 with acid. Add to this solution a quantity of sodium phosphate buffer salt to make the final concentration 0.1-M phosphate when the total volume is diluted to 100 ml. Stir and readjust the pH to 7.2 using acid or base. The final solution is 1.52-M diamine, 0.1-M sodium phosphate, pH 7.2. Allow the solution to come to room temperature before continuing.

3. Add the washed periodate-oxidized agarose to the diamine solution and mix to resuspend the gel.

4. Add 0.63 g of sodium cyanoborohydride (toxic!) to the gel slurry and mix for 2 to 4 h. Mixing may be carried out using an overhead paddle stirrer (not a stir bar) or in a sealed plastic container by end-over-end rocking. Avoid the use of sealed glass containers, because there is some gas evolution during the reductive amination process.

5. Extensively wash the diamine-agarose support with water, 1-M NaCl, and water to completely remove unreacted diamine and reaction byproducts. The support may be stored as a 50% slurry containing a preservative at 4°C until use.

(B) Coupling of a BOC–Amido–PEG–Amine Spacer to CDI-Activated Supports

The following protocol may be used to couple a bis-amino–PEGn compound that has one amino end blocked with a BOC protecting group. The support material used in this procedure must be able to withstand the depro-tection conditions for removing the BOC group using formic acid in solvent. Small samples of a given support should be tested for stability in a formic acid/DMAC solution before proceeding to use this procedure. Many crosslinked or polymeric supports should be able to tol-erate the deprotection step without difficulty.

1. In a fume hood, drain 10 ml of A CDI-activated support of excess acetone using a sintered glass filter funnel suspended in a suction filter flask (see Carbonyl Diimidazole (CDI) Activation in Section 2.1, this chapter). While pulling a gentle vacuum to remove the remaining excess acetone, break the support up into small pieces so that it resembles fluffy snow. Do not allow the support to dry. Remove the vacuum and resuspend the matrix in dry DMAC with mixing. Wash with DMAC to remove the last traces of acetone (at least 10 bed volumes). Drain to a moist cake.

2. Dissolve the blocked diamine compound BOC-N–amido–dPEG11–amine (Quanta Biodesign) in 10 ml of DMAC at a concentration of 12.8 mg/ml, which will equal a level of 20 μmol/ml gel in the reaction medium. Control of the final density of amines on

the support can be accomplished by adjusting the reaction concentration of the spacer. The optimal concentration of spacer to be used in the coupling reaction should be determined experimentally on small quantities of activated support to identify the best final affinity support performance in its intended application. Add an organic base to the ligand solution, such as DMAP, DIEA, or TEA, to make a final concentration of 2-mM.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for 1 to 2 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. Longer reaction times may be used if appropriate.

4. Excess CDI reactive groups on the support may be blocked by the addition of ethanolamine to the reaction slurry at a final concentration of 0.1-M. Continue to mix for 1 h at room temperature.

5. Transfer the gel slurry to a sintered glass filter in the fume hood that is suspended in a suction filter flask and wash extensively (at least 10 bed volumes) with solvent to remove the remaining ligand and reaction byproducts. Finally, drain the support of excess solvent by pulling a gentle vacuum on the filter flask while breaking up the support into small, finely divided pieces using a spatula, but be careful not to allow the matrix to dry out. Once the support is broken into small pieces, remove the vacuum and resuspend the gel in neat formic acid with mixing (caution: highly corrosive; use a fume hood and personal protective equipment to avoid contact or inhalation of vapors). Stir for 1 h at room temperature.

6. Wash the support with 2 bed volumes of formic acid and then wash extensively with DMAC to completely remove all traces of remaining acid. Once the acid has been thoroughly removed, the support should be washed into water by sequential washes using increasing concentrations of water in DMAC until 100% water is attained. Continue to wash the support with water until all traces of solvent have been removed. Additional washes with 1-M NaCl as well as low and high pH conditions may be done as appropriate. Finally, wash with water and store the amine-containing support as a 50% slurry in water containing a preservative at 4°C until used.

AMINO–CARBOXYLATE SPACERS

Spacer arms containing an amine group on one end and a carboxylic acid group on the other end are pop-ular choices for building many affinity supports. This type of spacer is typically used to modify a support to contain a terminal carboxylate for further coupling to amine-containing molecules. The spacer’s cross-bridge



can be designed to possess additional properties that may be beneficial to the final affinity support being cre-ated. For instance, short or long spacers can be chosen to modify the support, depending on the total length desired to tether a ligand away from the matrix surface. In addition, the spacer cross-bridge can be designed to be relatively hydrophobic (aliphatic) or hydrophilic (typically hetero-aliphatic construction).

The shortest amino–carboxylate spacer is β-alanine, which contains two central methylene groups with an amine on one end and a carboxylic acid group on the other end (3-aminopropanoic acid). This short spacer is used often in the design of modification and cross-linking agents, and it also can be used as a spacer on chromatography supports to provide a terminal car-boxylate for subsequent coupling reactions. β-Alanine is extremely water soluble and will not contribute any hydrophobicity to a support, which may occur with a longer aliphatic cross-bridge; however, the molecule also is quite short, which means it cannot be used to extend a ligand out from the matrix to facilitate greater binding accessibility for a target molecule. It is mainly used just to form a carboxylate on the support for fur-ther modification reactions or for facilitating ligand immobilization.

One of the most common, medium length, aliphatic amino–carboxylate spacers is 6-aminocaproic acid, which contains a six-atom methylene cross-bridge between the two functional groups. This compound has been widely used to modify amine-reactive supports to contain carboxylates for further immobilization of an amine-containing ligand. Usually, the use of a spacer is done to move the ligand away from the matrix to allow efficient docking of target molecules to bind with the ligand. For instance, Bansal et al. (2006) coupled p-aminobenzamidine by its amine to a monolithic cryo-gel containing aminocaproic acid spacers and used the resultant affinity support to isolate urokinase from cell culture broth of human kidney cells. The hydrophobic nature of the cross-bridge in 6-aminocaproic acid can

provide some enhancement of binding toward certain target molecules, depending on the circumstances of the interaction. The spacer has found use in other areas of bioconjugation as well, including its incorporation into the very popular biotinylation reagent, NHS–LC-biotin (see Chapter 11, Section 6.2). The 6-aminocaproic acid spacer has also been used to modify a matrix to create an NHS ester, amine-reactive support by forming the active ester on the terminal carboxylate after the spacer has been attached (Wilchek and Miron, 1987).

The spacer 6-aminocaproic acid is an inexpen-sive compound that has very good water solubility (50 mg/ml). It can be dissolved in aqueous buffers with-out significantly effecting the pH of the solution (unlike the diamine compounds described previously). The compound is an analog to the amino acid lysine and has antifibrinolytic properties in vivo by inhibiting plas-minogen. The coupling of 6-aminocaproic acid to an aldehyde-containing support using reductive amination is illustrated in Figure 15.112. Other amine-reactive sup-ports can be used in a similar process to create a carbox-ylate-terminal spacer.

COUPLING 6-AMINOCAPROIC ACID TO AN AMINE-REACTIVE SUPPORT The following protocol describes the modification of an aldehyde-containing support, such as periodate-oxidized agarose (see earlier section, this chapter) with 6-aminocaproic acid to provide a car-boxylate derivative for subsequent immobilization of amine-containing ligands. The subsequent immobiliza-tion of an affinity ligand to form an amide bond may be performed using the methods previously described in this chapter, such as a carbodiimide-mediated coupling reaction (using EDC) or through NHS ester formation.

1. Wash 100 ml of periodate-oxidized agarose (or another aldehyde-containing support) with water to remove storage solutions and then into coupling buffer (0.1-M sodium phosphate, pH 7.2). Washing steps may be performed with a sintered glass filter


O

H

+

NaCNBH3


H2N

6-Aminocaproic acid

HN

OH

O

OH

O

FIGURE 15.112 Coupling of 6-aminocaproic acid to an aldehyde-containing support via reductive amination.



funnel suspended in a vacuum filter flask. Wash with at least several bed volumes for each wash step. Drain to a moist cake.

2. In the fume hood, add 10 g of 6-aminocaproic acid (Acros, Aldrich) to 100 ml of coupling buffer. Stir the solution to dissolve and readjust the pH to 7.2 using acid or base, if necessary. The final solution is 0.76-M aminocaproic acid, 0.1-M sodium phosphate, pH 7.2. Lower concentrations of 6-aminocaproic acid may be used for this reaction to create a lower density of carboxylates on the surface of the support. Some experimentation may have to be carried out to determine the optimal level of carboxylates for the intended affinity chromatography application.

3. Add the washed periodate-oxidized agarose to the aminocaproic acid solution and mix to resuspend the gel.

4. In a fume hood, add 0.63 g of sodium cyanoborohydride (toxic!) to the gel slurry and mix for 2–4 h. Mixing may be performed using an overhead paddle stirrer (not a stir bar) or in a sealed plastic container by end-over-end rocking. Avoid the use of sealed glass containers, because there is some gas evolution during the reductive amination process.

5. Extensively wash the aminocaproic acid-agarose support with water, 1-M NaCl, and water to completely remove unreacted reagent and reaction byproducts. The support may be stored as a 50% slurry containing a preservative at 4°C until use.

AMINO–PEGn–CARBOXYLATE SPACERS

A version of an amine-carboxylate spacer that has a PEG-based cross-bridge is available that makes the linker extremely hydrophilic (Thermo Fisher, Quanta Biodesign). These spacers are particularly advanta-geous for creating support derivatives that have very low nonspecific binding character, unlike aliphatic spacers that may increase the nonspecificity of a sup-port. PEG-based spacers can also increase the freedom of motion of tethered affinity ligands, thus making them more likely to be able to interact with a target molecule. A series of amino–PEGn–carboxylate com-pounds are available depending on the spacer need, ranging from PEG4 to PEG36 in cross-bridge length (Quanta). The shortest is an amino–PEG4–carboxylate that has a 16 atom spacer about 18 Å in length. A lon-ger length amino–PEG8–carboxylate compound has a 33.6-Å spacer and the amino–PEG12–carboxylate reagent contains a 46.8-Å tether. The longest compound in this series has an internal PEG36 chain with a linear cross-bridge of 132.7 Å, making it perhaps the longest discrete spacer available for modification of chromatog-raphy supports. The largest spacer arm is actually lon-ger than the average diameter of a typical IgG antibody

molecule (~110 Å), which ensures that any affinity ligand tethered at the end of it will be accessible to vir-tually any docking protein or biomolecule that may bind to it (Figure 15.113).

Amino–PEGn–carboxylate spacers can be reacted with an amine-reactive support to give a covalent link-age forming an amide or secondary amine bond with the surface. The terminal carboxylate then can be used to build another reactive group for subsequent coupling of an affinity ligand, such as in the creation of an NHS ester or a hydrazide group (see previous sections, this chapter). The carboxylate may also be used to directly immobilize an amine-containing molecule through the use of a carbodiimide-mediated reaction sequence, which ultimately will form an amide bond with the ligand (see previous section, this chapter, on EDC cou-pling). A possible reaction sequence using these spacers is illustrated in Figure 15.114. The use of these long PEG spacer molecules will create an extremely hydrophilic layer on any solid-phase surface, which will lower non-specific binding character and maximize the purity of any target molecule captured.

Any of the amine-reactive immobilization methods described previously in this chapter can be used to cou-ple an amino–PEGn–carboxylate spacer to a support. In the following protocol examples, first a DSC-activated hydroxylic support is used in nonaqueous solvent to couple the amino–PEGn–carboxylate spacer to form an amide bond and in the second example a periodate-oxi-dized agarose support containing aldehyde groups is used to couple the spacer using reductive amination to form a secondary amine bond. Other reactive supports may be used by following the recommended coupling protocols found in previous sections.

COUPLING AMINO–PEGn–CARBOXYLATE SPACERS (A) Coupling Amino–PEG8–Carboxylate to a DSC-Activated Hydroxyl-Support1. In a fume hood, drain 10 ml of A DSC-activated

support of excess acetone using a sintered glass filter funnel suspended in a suction filter flask. While pulling a gentle vacuum to remove the remaining excess acetone, break the support up into small pieces so that it resembles fluffy snow. Do not allow the support to dry. Remove the vacuum and resuspend the matrix in dry DMAC with mixing. Wash with DMAC to remove the last traces of acetone (at least 10 bed volumes). Drain to a moist cake.

2. Dissolve the amino-PEG8-carboxylate spacer compound (Quanta BioDesign or Thermo Fisher) in 10 ml of DMAC at a concentration of 8.83 mg/ml, which translates into a level of 20 μmol/ml gel in the reaction medium. Control of the final density of amines on the support can be accomplished by adjusting the reaction concentration of the



spacer. The optimal concentration of spacer to be used in the coupling reaction should be determined experimentally on small quantities of activated support to identify the best final affinity support performance in its intended application. Add an organic base to the ligand solution, such as DMAP, DIEA, or TEA, to make a final concentration of 2-mM.

3. Add the activated wet gel cake to the ligand solution with stirring to fully resuspend the gel. Mix the reaction slurry for 1 h using an overhead paddle stirrer or by end-over-end rocking in a sealed container. Longer reaction times may be done if appropriate.

4. Excess NHS carbonate reactive groups on the support may be blocked by the addition of ethanolamine to the reaction slurry at a final concentration of 0.1-M. Continue to mix for 1 h at room temperature.

5. In the fume hood, transfer the gel slurry to a sintered glass filter that is suspended in a suction filter flask and wash extensively (at least 10 bed volumes) with solvent to remove the remaining ligand and reaction byproducts. Finally, drain the support of excess solvent by pulling a gentle vacuum on the filter flask while breaking up the support into small, finely divided pieces using a spatula, but be careful not to allow the matrix to dry out. Once the support is divided into small pieces, remove the vacuum and resuspend the gel in water with mixing. Continue to wash the support with water until all traces of solvent have been removed. Additional washes with 1-M NaCl as well as low and high pH conditions may be done as appropriate. Finally, wash with water and store the carboxylate-containing support as a 50% slurry in water containing a preservative at 4°C until used.

H2NO

OO

O

H2NO

OO

OO

OO

O

H2NO

OO

OO

OO

OO

O

O

O

OO

OO

OO

OO O

O

OOO

OO

OO

OO

OH2N

OO

O

OH

O

OHO

O

OH

O

OH

O

Amino–PEG4–carboxylate




FIGURE 15.113 Examples of amino–PEGn–carboxylate spacers.



(B) Coupling Amino–PEG8–Carboxylate to a Periodate-Oxidized Agarose Support1. Wash 10 ml of periodate-oxidized agarose (or another

aldehyde-containing support, such as a glycidol/periodate-treated polymeric support) with water to remove storage solutions and then into coupling buffer (0.1-M sodium phosphate, pH 7.2). The washing steps may be done with a sintered glass filter funnel suspended in a vacuum filter flask. Wash with at least several bed volumes for each wash step. Drain to a moist cake.

2. In the fume hood, add 176 mg of amino–PEG8–carboxylate spacer compound (Quanta BioDesign or Thermo Fisher) to 10 ml of coupling buffer. Stir the solution to dissolve and readjust the pH to 7.2 using acid or base, if necessary. The final solution is at a concentration of 40 μmol/ml of the spacer in 0.1-M sodium phosphate, pH 7.2. Higher or lower concentrations of amino–PEG8–carboxylate may be used for this reaction to create custom densities of carboxylates on the surface of the support. Some experimentation may have to be done to determine

the optimal level of carboxylates for the intended affinity chromatography application.

3. Add the washed periodate-oxidized agarose to the amino–PEG8–carboxylate solution and mix to resuspend the gel.

4. Add 63 mg of sodium cyanoborohydride (toxic!) to the gel slurry and mix for 2 to 4 h. Mixing may be performed using an overhead paddle stirrer (not a stir bar) or in a sealed plastic container by end-over-end rocking. Avoid the use of sealed glass containers, because there is some gas evolution during the reductive amination process.

5. Extensively wash the carboxylate-agarose support with water, 1-M NaCl, and water to completely remove unreacted reagent and reaction byproducts. The support may be stored as a 50% slurry containing a preservative at 4°C until use.

BIS-CARBOXYLATE SPACERS

bis-Carboxylate spacer compounds are useful in extending an amine-containing support and converting the amine to a terminal carboxylate for further coupling

H2NO

OO

O OH

O


O

CDI-activated support

O

N

N

+O

O

NH

OO

OO OH

O

Support containing terminal carboxylate

O

O

NH

OO

OO O

O

DSC inDMAC

N

O

O

NHS ester–activated support

O

O

NH

OO

OO

O

NH


Ligand coupling throughamide linkage

H2N

FIGURE 15.114 Coupling of an amino–PEGn–carboxylate to an amine-reactive support followed by NHS ester formation and the immobili-zation of an amine-containing ligand.



or immobilization reactions. Most often, a bis-carboxyl-ate spacer is added to a support through reaction of its corresponding cyclic anhydride with the amino groups on a matrix to create amide bond linkages. Ring open-ing of the anhydride forms the short spacer group as the other carboxylate is freed at the uncoupled end. Several anhydride compounds are commonly used to form these spacers on supports (Figure 15.115). Succinic anhydride is the shortest of these spacers, containing a 4-carbon ring structure, which when reacted with an amine-containing support provides a 2-atom methylene bridge between the two terminal carboxylates. Glutaric anhydride is a 5-carbon compound that when opened results in a 3-atom methylene bridge between the two terminal carboxylates. Diglycolic anhydride is the most hydrophilic in this series, which after ring open-ing contains a 5-atom spacer between the two terminal carboxylate groups and also has a central ether group, which adds water solubility to the compound. All three anhydride compounds may be used in similar reac-tion schemes to modify an amine-containing support to contain terminal carboxylates. The reaction of succinic anhydride with an amine-containing support is illus-trated in Figure 15.116.

The following protocol describes the modification of an amine-containing support with succinic anhydride to form a carboxylate-containing support for further ligand immobilization reactions. The other anhydride compounds may be used similarly.

Coupling Succinic Anhydride to Amine-Containing Supports 1. Wash 100 ml of an amine-containing support with

water to remove storage solutions and then wash with several bed volumes of reaction buffer (1.0-M sodium bicarbonate, pH 8.0). Finally, suspend the

support in an equal volume of reaction buffer. Other reaction buffers and pH conditions may be used over the range of about pH 6 to pH 9 with success. Some protocols just use water, but maintain the pH at the desired value with periodic titration with base (NaOH at 50%).

2. Stir the gel slurry using an overhead paddle stirrer and slowly add 10 g of succinic anhydride. Initially, the anhydride will not be fully soluble in the reaction mixture, but it will go into solution as it reacts and hydrolyzes in the buffered solution.

3. React with constant stirring for 1 h at room temperature.

4. Wash the succinylated support with reaction buffer, water, 1-M NaCl, and again with water to completely remove unreacted succinic anhydride or succinic acid. The carboxylic acid-containing support may be stored as a 50% slurry containing a preservative at 4°C until use.

Other anhydride compounds may be used in a simi-lar reaction protocol with success; however, adjust the mole quantity of anhydride addition to be equivalent to the succinic anhydride protocol.

THIOL–CARBOXYLATE SPACERS

Spacer arms containing a thiol group on one end and a carboxylate group on the other end have been used extensively for some applications involving immobilized affinity ligands. Most of these applications have involved the use of a solid phase that contains a metallic surface, either planar or particulate in nature. In this case, the thiol end of the spacer is usually coupled to the metal by way of a dative bond, wherein the unshared pair of electrons on the sulfur atom are shared with the metal. The application of these spacers to modify metallic mic-roparticles or nanoparticles is described in Chapter 14. Although chromatography supports typically do not use metallic solid phases, in certain instances it may be advantageous to use a thiol–carboxylate spacer to

OO O

Succinic anhydride

OO O

Glutaric anhydride

O

OO O

Diglycolic anhydride

FIGURE 15.115 The structures of common anhydride com-pounds used to create terminal carboxylate groups on amine-contain-ing supports.

OO O

Succinic anhydride

+


NH2

HN

OOH

O

Carboxylate-containing support

FIGURE 15.116 Reaction of succinic anhydride with an amine-containing support.



modify an amine-containing matrix with the carboxyl-ate end, and thus create a thiol group on a support for subsequent immobilization of thiol-containing ligands. Terminal thiols created in this manner on a support can be activated to couple ligands through disulfide bonds, which then can be reversed to subsequently release inter-acting molecules (see previous section on thiol-reactive immobilization methods, this chapter). Of course, it also is possible to use a thiol–carboxylate spacer to modify a thiol-reactive support to create a terminal carboxylate, but this probably is less frequently done than coupling the spacer in the reverse way.

A number of thiol–carboxylate spacer arms are avail-able to modify a support, including molecules that contain aliphatic cross-bridges as well as the more hydrophilic PEG-based linkers. The shortest thiol–car-boxylate spacer is thioglycolic acid, which is 2-mercap-toacetic acid (note that this compound is not thioacetic acid, which is the acetylated thiol in a thioester form). An analog to thioglycolic acid is SATA (Chapter 2, Section 4.1), which has a protected thiol at one end and an NHS ester at the other end of the acetate bridge. SATA, or the similar compound SATP which is one car-bon atom longer, can provide rapid modification of an amine-containing support to convert it to a protected thiol-containing support. The NHS ester end will react with the amines on the support to result in an amide bond, and the protected thiol end is stable in this form until needed for immobilization of ligands. The thiol can be deprotected with hydroxylamine under alkaline conditions to reveal the thiol. These short compounds can be useful if an amine-containing support already has a long spacer arm present and all that is desired is to convert a terminal amine into a thiol group.

There also are longer thiol–carboxylate spacers avail-able to form tethers of greater length on supports for the coupling of affinity ligands. Both hydrophobic, aliphatic spacer molecules can be used as well as hydrophilic, PEG-based spacers. A PEG-containing spacer will pro-vide greater hydrophilicity within the support and have a tendency to decrease the nonspecific binding poten-tial of biomolecules. There is a wide selection of differ-ent spacer lengths available for these compounds with PEG groups ranging from PEG4 to PEG20 in size (Quanta BioDesign, Thermo Fisher). However, the use of a thiol-carboxylate spacer without having the thiol group pro-tected can be problematic. The reactions that have to occur to couple the carboxylate end to an amine-contain-ing support are unfortunately interfered with by the thiol end. If an active ester is formed on the carboxylate to couple with the amines on the support without protect-ing the thiol, then the ester can potentially react with the thiol groups to form thioester bonds, causing polymer-ization of the reagent in solution. In addition, if the car-bodiimide EDC is used to attach the carboxylate to the

amino groups, then EDC can also react with the thiol end to form an irreversible complex. Fortunately, thiol-pro-tected versions of these reagents are available that allow these reactions to occur without interference. These com-pounds are SATA-like in structure, having an NHS ester on one end and a protected (acetylated) thiol on the other end, but with the added feature of containing an inter-nal PEG cross-bridge to provide increased hydrophilicity over that of the aliphatic reagent. Figure 15.117 illustrates the major thiol-carboxylate reagents available for use as spacers in chromatography supports and Figure 15.118 shows the reactions involved with coupling one of these reagents to an amine-containing support.

Coupling NHS–PEG4–Thioacetyl to an Amine-Containing Support

The following protocol describes the use of a NHS–PEG4–thioacetyl compound in the modification of an amine-containing support to provide an altera-tion that terminates in a protected thiol group. The reagents SAT(PEG)4 from Thermo and dPEG4-SATA from Quanta BioDesign are identical structurally and can be used in this method with success. This method describes the reaction of the NHS ester end with the amine groups on the support in a nonaqueous envi-ronment to eliminate the hydrolysis of the ester during coupling. All operations with solvent should be per-formed in a fume hood using the appropriate personal protective equipment. The reaction may also be car-ried out in an aqueous buffer (e.g., 0.1-M sodium phos-phate, pH 7.2), but the amount of reagent added may have to be increased to account for some yield loss due to hydrolysis. The optimal density of protected thiols on the support should be investigated experimentally by assessing the performance of the ultimate affinity support in the intended application. Adjusting the con-centration of the reagent in the reaction will control the density of the protected thiols after the reaction is com-plete, and so will adjusting the initial concentration or density of amines on the support.

1. Wash 10 ml of an amine-containing support with water to remove storage solution and preservatives. Then wash the support into DMAC by washing with sequentially increasing concentrations of solvent-in-water until 100% DMAC is used. Continue to wash with DMAC for at least 10 to 20 bed volumes to completely remove the last traces of water. Drain the support to a wet cake, but do not allow it to dry.

2. Dissolve 421 mg of the NHS–PEG4–thioacetyl compound in 10 ml of DMAC. This will result in 100 μmoles of reagent per milliliter of amine-containing gel within the reaction mixture.

3. Add the washed gel to the reagent solution with mixing to resuspend the matrix. An organic base



may be added to the mixture to accelerate the reaction, such as DMAP, DIEA, or TEA, to make a final concentration of 2-mM. React for 1 h at room temperature with constant mixing. The mixing may be accomplished by use of a paddle and overhead stir motor or by end-over-end rocking in a sealed container.

4. Wash the support with DMAC to remove excess reagent (at least 10 column volumes) and then wash

sequentially back into water by using increasing concentrations of water-in-solvent until 100% water is used. Continue to wash with water for 10 to 20 bed volumes to completely remove the solvent. The protected thiol-containing support may be stored at 4°C as a 50% slurry containing a preservative until use. As needed, the gel can be treated to deprotect the thiol groups prior to immobilizing a ligand or doing further reactions.

HSO

OHThioglycolic acid

N

O

O

O

OS

OSATA

N

O

OO S

O O

SATP

OO

OO

ON

O

O

OS CH3

O

SAT–PEG4

NH

OO S

N-Acetyl homocysteinethiolactone

O

OHHS

8-Mercaptooctanoic acid

OO

OO

ON

O

O

OO CH3

OO

OO

S

SAT–PEG8

OO

OO

ON

O

O

OO

OO

OO

O

OO

SH3C

O

SAT–PEG12

OO

OO

ON

O

O

OO

OO

OO

O

OO

O

SAT–PEG16

OO

OSH3C

O

OO

OO

ON

O

O

OO

OO

OO

O

OO

O

SAT–PEG20

OO

OO

OO

O

CH3O

S

FIGURE 15.117 Thiol–carboxylate reagents for spacer arm applications.



Deprotection of the Thioacetyl-Containing Support 1. Prepare 20 ml (or 2 times the amount of protected

thiol-containing resin to deprotect) of an aqueous solution containing 0.5-M hydroxylamine in 0.1-M sodium phosphate, pH 7.2, containing 25-mM EDTA.

2. Wash 10 ml of the protected thiol resin with several bed volumes of water to remove storage solution and then slowly wash with 10 ml of the hydroxylamine solution. Drain to a wet cake.

3. Add the washed support to the remaining 10 ml of hydroxylamine solution and mix to resuspend the gel. React for 2 h at room temperature with constant mixing to remove the acetyl protecting groups.

4. Wash the deprotected support extensively with water to completely remove the hydroxylamine solution and reaction byproducts. The thiol-containing matrix should be used immediately to immobilize a ligand or in another reaction to couple with the free –SH group. Avoid storage of the support in this form, because thiol oxidation likely will take place rapidly and degrade the amount of thiols available. EDTA (25-mM) may be added to the wash solutions or reaction buffers to prevent metal-catalyzed oxidation.

OO

OO

ON

O

O

OS CH3

O

SAT–PEG4

+


NH2

HN

O

OO

OO

S CH3

OIntermediate protected thiol

HN

O

OO

OO

SH

Hydroxylamine

CH3

O

HN

HO

Support containing free thiols

+

FIGURE 15.118 Coupling of a NHS–PEGn–thioacetyl reagent to an amine-containing support with subsequent deprotection of the thiol using hydroxylamine.

bioconjugate techniques || immobilization of ligands on chromatography supports

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