Hermanson G. Bioconjugate Techniques, Second Edition

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590 14. Microparticles and Nanoparticles

the polymeric diversity of these particles, even of those that consist of a styrene base, it is best

to identify the particle type by its exact chemical composition.

Polymeric particles also can be further diversiﬁ ed by their surface compositions. Even tra-

ditional polystyrene particles contain additional groups on their surfaces due to the poly-

merization reaction used to create them. For instance, nearly all such particles contain some

negatively charged sulfonate groups, which are the result of the catalyst, persulfate used to ini-

tiate the polymerization process. In addition, many particles contain non-covalently adsorbed

detergent molecules that can add amphipathic character to the particle surface, depending on

the type of detergent used during the polymerization reaction. Detergents can be removed by

extensive washing after the manufacturing process, but this may be a source of variability from

manufacturer to manufacturer.

Another way of altering surface characteristics is through the purposeful addition of second-

ary polymers onto a polymeric core by graft copolymerization, adsorptive coating, or through

covalent attachment of another polymer type. Adding hydrophilic coatings onto hydrophobic

particle cores often is done to promote biocompatibility and limit nonspeciﬁ c interactions with

proteins or other biomolecules. Examples of hydrophilic coat polymers that have been used for

this purpose include poly(ethylene glycol) (PEG), poly(vinyl alcohol) (PVA), poly(acrylic acid),

poly(methacrylic acid), poly(acrylamide), and poly(vinyl pyrrolidone). Hydrophilic modiﬁ ca-

tion of hydrophobic particle cores can aid in blocking the nonspeciﬁ c binding potential that the

core may have toward proteins or other biological molecules. It also can alleviate the tendency

of particles to aggregate in aqueous solution by creating a hydrated shell around each particle,

which helps keep them separated and in suspension. By example, Harper et al. (1995) describe

the copolymerization of a poly(ethylene oxide) methacrylate monomer (PEG-type) to create

polystyrene particles with hydrophilic surfaces for the attachment of cells.

Covalent attachment of hydrophilic spacer arms to particles is another way of masking the

hydrophobic character of the underlying surface while making a particle more biocompatible.

Spacers of this type are typically short organic molecules containing a number of polar groups

along the chain, which are able to interact with the surrounding aqueous environment and limit

nonspeciﬁ c protein adsorption on the particles. A common spacer arm construction consists of a

PEG chain that also may include a short hydrophobic straight chain linker, which attaches to the

particle surface. To provide an attachment site for afﬁ nity ligands to the spacer, a functional group

can be included on the outer end of the PEG polymer, such as a carboxylate group. These func-

tional groups provide sites for covalent coupling of afﬁ nity ligands, including proteins, while the

PEG chain creates a hydrophilic “lawn” to limit nonspeciﬁ c interactions. The best conﬁ guration of

this spacer type is to have about 10 percent of the spacers terminate in a carboxylate group while

the rest of them terminate in a PEG hydroxyl or a PEG methyl ether group. This design has been

used with success with other surface types, such as gold nanoparticles or planar surfaces (Prime and

Whitesides, 1991), and it can be adapted to polymer particles with little difﬁ culty ( Figure 14.5 ).

4.1. Passive Adsorption

One of the simplest methods of attaching biomolecules to hydrophobic polymeric particles is

the use of passive adsorption. Some of the earliest examples related to the use of particles in

immunoassays include the use of non-covalently adsorbed antibody or antigen onto latex micro-

spheres. Protein adsorption onto hydrophobic particles takes place through strong interactions

of non-polar or aromatic amino acid residues with the surface polymer chains on the particles

with concomitant exclusion of water molecules. Since proteins usually contain hydrophobic core

structures with predominately hydrophilic surfaces, their interaction with hydrophobic parti-

cles must involve signiﬁ cant conformational changes to create large-scale hydrophobic contacts.

These conformational changes often result in complete denaturation of the ﬁ rst protein layer to

be adsorbed onto the particles. Subsequent protein molecules which bind and add to this ini-

tial layer are adsorbed through protein–protein binding or aggregation events, which ultimately

Figure 14.5 A method of making particles biocompatible includes the use of PEG-based spacers. A lawn of

mPEG molecules in interspersed with some longer PEG chains that terminate in carboxylate groups for coupling

amine-containing molecules. The result is an extremely hydrophilic surface with low nonspeciﬁ c binding.

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592 14. Microparticles and Nanoparticles

result in the formation of protein clusters in which the outer layers of protein have more of a

native conformation and activity (Butler, 2000a, b).

Due to this tendency of proteins to form multilayer coatings on hydrophobic surfaces, the

amount of excess protein beyond the particle saturation point (theoretical monolayer density

for the protein type) added during adsorption processes should be limited. If extremely large

excesses of protein are added to particles, highly unstable constructs may result which may con-

tinually leach off loosely adsorbed protein. Most protocols recommend a protein concentration

of about 3–10  excess of protein over the calculated monolayer concentration for the particles

used. An estimate of the maximal amount of a given protein that can bind as a monolayer to a

particle surface can be determined by the fact that bovine serum albumin (BSA) (MW 67 KDa)

has an adsorption capacity of about 3 mg/m

of particle surface. A larger protein like an anti-

body (IgG; MW 150 KDa) has a maximal adsorption density of about 2.5 mg/m

of particle

surface. To translate this into a real-world example, 1 g of 1 m polystyrene microspheres can

adsorb about 18 mg BSA or about 15 mg IgG (Cantarero et al., 1980). Knowledge of a parti-

cle’s surface area per gram will permit reasonable estimates of maximal adsorption capacity for

a given protein relative to the size of an antibody or albumin molecule.

Passive adsorption has been studied extensively as it relates to the immobilization of anti-

body molecules onto microplates (polystyrene) or microspheres (hydrophobic beads of various

compositions). Although the denaturing effect of passive adsorption of proteins onto surfaces

was known since 1956 when Bull studied the adsorption of albumin onto glass, it was not

widely recognized as being detrimental until a number of more recent studies were published.

See in particular Butler et al. (1992) which investigates the physical and functional behavior of

capture antibodies adsorbed on polystyrene, Butler (2000a) which provides a summary of the

effects of the adsorption of various proteins on hydrophobic surfaces, Cantarero et al. (1980)

which discusses the adsorptive characteristics of proteins for polystyrene surfaces, and Butler

et al. (1997) which compares the effect of passive adsorption on the antigen speciﬁ city of

immobilized antibody molecules.

The detrimental effects on biomolecules as a result of passive adsorption onto hydrophobic

surfaces can be dramatic. Butler et al. (1993) determined that over 90 percent of monoclonal

antibodies and about 75 percent of polyclonal antibodies against ﬂ uorescein were denatured

upon adsorption onto polystyrene surfaces. In addition, Bagchi and Birnbuam (1981) observed

that IgG was optimally adsorbed to poly(vinyltoluene) particles at a pH value near the anti-

body’s pI, indicating that the interactions were entirely hydrophobic in nature and didn ’t

depend on charge interactions. Once the antibody was adsorbed, however, it was tightly bound

to the surface, provided that the pH was maintained at the initial adsorption point. If any pH

cycling was done, such as may occur with some wash steps, the adsorbed antibody had a ten-

dency to leach off the surface, demonstrating the non-covalent nature of the interaction.

Protein adsorption to hydrophobic polymer particles thus should be done at or near the isoelec-

tric point of the particular protein to assure the highest density of protein is attached to the parti-

cles. Proteins at their isoelectric point exist in the most collapsed conformation possible, because

charge repulsion effects don ’t play as signiﬁ cant a role on overall globular structure. Upon adsorp-

tion under isoelectric conditions, each protein then can bind to the polymer surface at maximal

density. Therefore, most suggested buffer cocktails used for passive adsorption recommend pH

conditions somewhere in the pI range of the protein. Since many proteins have a range of different

biological modiﬁ cations (post-translational) resulting in a range of pI values, some optimization

may need to be done to determine the best pH conditions to use for an adsorption process.

Another factor to consider is the amount of protein available for adsorption. If expensive

antibody in very small amounts is used to passively adsorb to polymeric particles, often there

is not enough antibody available to saturate the particle surface. In this case, another carrier

or blocking protein may be added to the mixture to take up otherwise unoccupied sites on the

particle surface. This will prevent particle aggregation that may occur if low concentrations of

protein are used in the adsorption process. Carrier proteins frequently used for this purpose

include albumin or polyclonal IgG pools, such as bovine gamma globulin. Other standard pro-

tein blocking agents also may be used for blending with the desired antibody, such as casein,

non-fat dried milk proteins, ﬁ sh serum, etc.

The polymeric particle composition also plays a signiﬁ cant role in the amount of protein

adsorbed and its stability and activity after immobilization. In general, copolymer blends

in which a hydrophobic monomer is polymerized with a charged monomer (such as acrylic

acid or methacrylate) create a surface that has both hydrophobic character along with areas

of charge. Proteins adsorbed onto these copolymer particles interact with the surface through

hydrophobic interactions and positive–negative charge attraction. The result is less protein

denaturation upon adsorption and better retention of activity after immobilization. A compari-

son of protein adsorption onto pure polystyrene and several other copolymer particle types by

Bale et al. (1989) resulted in the following order of binding afﬁ nity:

Polystyrene  polystyrene/PMMA copolymer  PMMA  polystyrene/polyacrylic acid copolymer

Thus, pure hydrophobic polymer compositions adsorb proteins very strongly, but have the

greatest tendency to denature protein at the initial protein–surface interface. Copolymers con-

taining some polarity or negative charge character bind protein less avidly, but result in better

retention of activity. The inclusion of some polar hydrophilic groups within an overall hydro-

phobic surface structure has been used with success for polystyrene microplates, as most of the

so-called high binding plates contain a population of polar constituents on the well surfaces,

allowing both hydrophobic and charge or dipole interactions to occur.

The following protocol for passive adsorption is based on methods reported for use with

hydrophobic polymeric particles, such as polystyrene latex beads or copolymers of the same.

Other polymer particle types also may be used in this process, provided they have the necessary

hydrophobic character to promote adsorption. For particular proteins, conditions may need to

be optimized to take into consideration maximal protein stability and activity after adsorption.

Some proteins may undergo extensive denaturation after immobilization onto hydrophobic

surfaces; therefore, covalent methods of coupling onto more hydrophilic particle surfaces may

be a better choice for maintaining native protein structure and long-term stability.

Protocol

1. Dilute the particles to a mass concentration of 10 mg/ml (1 percent) in coating buffer at

a pH near the pI of the protein being adsorbed. If particle aggregation becomes a prob-

lem, the initial particle concentration may be reduced to 5 mg/ml. Some typical coating

buffer suggestions include: (a) 10 mM sodium phosphate, 0.15 M NaCl, pH 7.4 (PBS);

(b) 50 mM sodium borate, pH 8.5; (c) 50 mM sodium acetate, pH 3.6–5.6; (d) 25 mM

MES [2-( N-morpholino)ethane sulfonic acid], pH 6.1; or (e) 50 mM sodium bicarbonate,

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594 14. Microparticles and Nanoparticles

pH 8.5–9.5. NaCl may be added to any of these buffers at a ﬁ nal concentration of

0.15 M to promote protein stability, if required. However, avoid the use of detergents or

chaotropic agents, as these will compete with protein adsorption or reduce the strength

of hydrophobic interactions, thus limiting the yield.

2. Dissolve the protein to be adsorbed in coating buffer at a concentration such that when

the solution is mixed with the particle suspension, the appropriate protein concentration

is reached to obtain an excess of about 3–10  over the maximal monolayer density for

that protein on the particles (see previous discussion). Note: A good strategy is to perform

a protein titration study, wherein a range of protein concentrations are tried with a given

particle quantity. For instance, using a 300 nm latex particle, a suitable set of trial experi-

ments would employ protein concentrations in the range of about 10–200 g protein/mg

particles. Plotting the amount of protein charged to the coating reaction versus the amount

bound will provide data to identify the optimal protein concentration to be used.

3. With rapid mixing, add protein solution to the particle suspension. For small volumes,

mixing can be done by pulling the solution up and down in a pipette tip or by vortexing.

For larger volumes, a stir bar or paddle may be used. Some protocols recommend adding

the particles to the protein solution to obtain the best initial mixing rate, thus assuring

even particle coating throughout the particle suspension. The key is rapid and efﬁ cient

mixing to create a homogeneous mixture of protein and particles, so adsorption can occur

uniformly on each particle.

4. Mix the suspension for 1 hour at room temperature.

5. Remove excess protein by centrifugation or tangential ﬂ ow ﬁ ltration. Many particles may

be pelletted successfully using a tabletop centrifuge, especially if the particle size is above

about 150 nm in diameter. If particle clumping is a problem upon pelletting or the particle

size is too small to be effectively pelletted, then ﬁ ltration is the better alternative. Centrifuge

and wash the particles at least twice with coating buffer to assure complete removal of

non-bound protein. Complete re-suspension of the particles may be accomplished by the

use of sonication, especially through the use of a sonic probe dipped into the solution.

6. Resuspend the particles to a ﬁ nal concentration of 1 percent in coating buffer containing

a preservative. Avoid changing the composition of the storage buffer from that used to

coat the protein, as any pH or compositional changes often result in elution of some of

the adsorbed protein.

7. Determine the amount of adsorbed protein on the particles by using a suitable protein

assay technique, such as the bifunctional chelating agents (BCA) Protein Assay (Thermo

Fisher).

4.2. Covalent Coupling to Polymeric Particles

Many particle types contain functional groups that are built into the polymer backbone and dis-

played on their surface. The quantity of these groups can vary widely depending on the type and

ratios of monomers used in the polymerization process or the degree of secondary surface modi-

ﬁ cations that have been done. Some common particle functionalities are shown in Figure 14.6 .

Many of these functionalized particles can be used to couple covalently biomolecules through the

appropriate reaction conditions (Illum and Jones, 1985; Arshady, 1993). For each type of particle,

manufacturers may offer several different densities of functional groups for different applications.

4.3. Coupling to Carboxylate Particles

One of the more common particle types is the carboxylate particle, which typically is created

by copolymerization or grafting with acrylic acid or methacrylic acid monomers. However, the

characteristics of carboxylate particles can vary between manufacturers and particle sources

due to the differences in surface density of carboxylate groups present. A measurement of

chemical density on particle surfaces is called the “parking area ”, and it refers to the average

area in Å

occupied by each functional group. A brief survey of carboxylate particles from

different manufacturers indicates that the average carboxylate parking area can vary widely

from about 10 Å

(1 nm

) to over 125 Å

(12.5 nm

). One carboxylate group every 125 Å

equivalent to about one carboxylate for every square having dimensions of 3.5 nm  3.5 nm,

or the space that might be occupied by one small protein on the particle surface ( note: albumin

(67 kDa) has an effective molecular diameter of about 7 nm, while an IgG molecule (150 kDa)

has a diameter of about 11 nm). At a density of one carboxylate every 125 Å

, there still may

be considerable polymer surface exposed, which could be a source for nonspeciﬁ c binding,

especially if the underlying polymer structure has hydrophobic character. A protein coupled to

Figure 14.6 Common functional groups or reactive groups on particles that provide the ability to couple pro-

teins or other ligands.

4. Polymeric Microspheres and Nanospheres 595

596 14. Microparticles and Nanoparticles

a carboxylate on such a particle could unfold through interactions with the exposed hydropho-

bic polymer surface and become denatured. Conversely, at the high end of carboxylate density,

there would be about one carboxylate present for every 1 nm

of particle surface. For the cova-

lent coupling of proteins, this high-density surface would provide more reactive sites for con-

jugation, while effectively masking the underlying polymer surface with negative charges. This

helps to maintain particle suspension through negative charge repulsion and prevents nonspe-

ciﬁ c binding or protein denaturation.

Carboxylate particles can be activated by a number of strategies, which yield reactive inter-

mediates capable of coupling with nucleophiles in proteins and other biomolecules. Figure 14.7

illustrates some of the reactions that can be used for coupling amine-containing molecules to

carboxylate particles, all of which form stable amide linkages with the particle. The water-solu-

ble reactions can be done in entirely aqueous conditions, both for activation and coupling. For

some reactions, however, the activation process must be done under nonaqueous conditions to

prevent hydrolysis of the activator or active intermediate. Activation in organic solvent is an

option for particles that remain stable in nonaqueous conditions. Some polymer particles may

unacceptably swell in organic solvent and they potentially could be damaged by such treatment.

By far the most common reaction strategy for coupling proteins and other amine-contain-

ing molecules to carboxylate particles is through an aqueous, carbodiimide-mediated process

using EDC (1-ethyl-3-(3-dimethylaminopropyl)carbodiimide), either in a single-step coupling

reaction or in a two-step reaction that employs the addition of N-hydroxysuccinimide (NHS)

or sulfo-NHS (see Chapter 3, Section 1.2 for additional information). Of all the crosslinking

methods used in bioresearch applications today, this relatively simple coupling process is the

most frequent conjugation reaction done with proteins. Using this reaction, carboxylate parti-

cles ﬁ rst are activated with the water-soluble carbodiimide EDC to create an intermediate ester.

This ester is reactive directly with amines on proteins, but it also can be used with the addition

of NHS or sulfo-NHS, which results in the formation of another intermediate, the NHS ester

or sulfo-NHS ester. The formation of this second ester results in a more stable intermediate in

aqueous solution than the one formed with EDC, therefore the secondary coupling reaction

with proteins proceeds with higher yields than with the use of EDC alone ( Figure 14.7 ). In

addition, by forming the secondary (sulfo)NHS ester, excess EDC can be removed from the par-

ticles before adding protein, thus preventing carbodiimide-mediated protein polymerization due

to the presence of both amines and carboxylates on most proteins (Staros, 1982; Borque et al. ,

1994; Bonﬁ eld et al., 2005).

For particle conjugation, it is important to maintain a repulsive force between particles to

stabilize the colloidal suspension, even during the activation and coupling reactions. For this

reason, the use of sulfo-NHS instead of NHS in this reaction results in an intermediate ester,

which is strongly negatively charged. The sulfonates on the sulfo-NHS esters actually are more

effective at keeping particles from aggregating than the original carboxylates; therefore, the

reactive intermediate particles easily can be puriﬁ ed from excess reactants and then mixed with

protein for coupling. If the non-sulfonated NHS is used in this reaction, the uncharged interme-

diate NHS ester may cause unacceptable particle aggregation, depending on the particle type,

although uncharged NHS has been used widely with success.

The following protocols for coupling amine-containing molecules to carboxylated particles

represent viable starting points for optimizing a method that works best for the particular mol-

ecule or protein being immobilized. The single-step method using EDC alone is appropriate for

use in coupling molecules having one or more amines present without any carboxylates. It works

especially well for small organic molecules, such as haptens, 5 -amino-modiﬁ ed oligonucleotides,

and amine-containing steroid derivatives (Hager, 1974; Quash et al., 1978; Nathan and Cohn,

1981; Fuller et al., 2006). If the molecule being coupled has both amines and carboxylates, such

as proteins, then it is best to use the two-step method, which eliminates excess EDC before the

addition of ligand, otherwise unacceptable ligand polymerization may take place.

Figure 14.7 Carboxylate particles can be coupled to amine-containing molecules using a number of reaction

strategies. The most frequently used method involves an aqueous two-step coupling process using EDC and

NHS or sulfo-NHS to form an amide bond with a protein or other molecules. It proceeds through an intermedi-

ate (sulfo)NHS ester, which has better reactivity for coupling amines than the initial EDC-reactive ester. Organic

solvent activation processes using NHS esters or acyl imidazole-reactive groups also can be used with solvent-

stable particles to result in the same product with an amine-containing molecule.

4. Polymeric Microspheres and Nanospheres 597

598 14. Microparticles and Nanoparticles

Single-Step EDC Coupling Protocol

1. Wash particles (e.g., 100 mg of 1 m carboxylated latex beads) into coupling buffer (i.e.,

50 mM MES, pH 6.0 or 50 mM sodium phosphate, pH 7.2; buffers with pH values from

pH 4.5–7.5 may be used with success; however, as the pH increases the reaction rate will

decrease). Suspend the particles in 5 ml coupling buffer. The addition of a dilute deter-

gent solution may be done to increase particle stability (e.g., ﬁ nal concentration of 0.01

percent sodium dodecyl sulfate (SDS)). Avoid the addition of any components containing

carboxylates or amines (such as acetate, glycine, Tris, imidazole, etc.). Also, avoid the

presence of thiols (e.g., dithiothreitol (DTT), 2-mercaptoethanol, etc.), as these will react

with EDC and effectively inactivate it.

2. Dissolve the amine-containing ligand to be coupled in 5 ml coupling buffer at a con-

centration sufﬁ cient to provide a 1- to 10-fold molar excess of ligand over the maximal

calculated carboxylate group concentration for the amount and type of beads used. For

particle manufacturers reporting a carboxylate concentration in meq/g, this is equivalent

to  mol/mg.

3. Combine the ligand solution with the particle suspension and mix thoroughly.

4. Add 100 mg EDC and mix to dissolve. To facilitate faster dissolution, EDC may be dis-

solved immediately before use as a concentrated stock solution in reaction buffer and

then an aliquot of this solution added to the particle suspension to obtain the correct

ﬁ nal concentration.

5. React at room temperature 2–4 hours with mixing.

6. Wash the beads and resuspend them in coupling buffer containing 30–40 mM of an

amine-containing hydrophilic quenching molecule to block excess reactive sites (i.e., eth-

anolamine or Tris).

7. Wash beads and store in an appropriate buffer containing a preservative.

Two-Step EDC/Sulfo-NHS Coupling Protocol

An alternative to this procedure was used by Kulin et al. (2002) for coupling antibodies to car-

boxylated microspheres, which provides different buffer conditions and activation with EDC

without the use of sulfo-NHS or NHS.

Activation

1. Wash particles (e.g., 100 mg of 1 m carboxylated latex beads) into coupling buffer

(50 mM MES, pH 6.0). Suspend in 5 ml coupling buffer. The addition of a dilute deter-

gent solution may be done to increase bead stability and prevent clumping (e.g., 0.01

percent SDS). Avoid the addition of any components containing carboxylates or amines

(such as acetate, glycine, Tris, imidazole, etc.). Also, avoid the presence of thiols (e.g.,

DTT, 2-mercaptoethanol, etc.), as these will react with EDC and effectively inactivate it.

2. Add 100 mg of EDC and 100 mg of sulfo-NHS. Mix to dissolve. To facilitate faster dis-

solution, EDC and sulfo-NHS may be dissolved immediately before use as a concentrated

stock solution in reaction buffer and then an aliquot of this solution added to the particle

suspension to obtain the correct ﬁ nal concentration.

3. React for 15 minutes at room temperature.

4. Quickly wash beads 2 times with coupling buffer using centrifugation and resuspend

using a sonic probe in 5 ml of the same buffer.

Coupling

5. Dissolve protein to be coupled in 5 ml coupling buffer at a concentration sufﬁ cient to provide

1- to 10-fold molar excess of ligand over the maximal calculated monolayer concentration

for the amount and type of beads used. For particle manufacturers reporting a carboxylate

concentration in meq/g, this is equivalent to mol/mg. The optimal protein concentration

should be optimized. Note: Too low a protein concentration may result in particle crosslinking.

For coupling of expensive antibodies that may not be available in enough quantity to reach

the optimal molar ratio on the particles, the addition of another protein (i.e., bovine gamma

globulin or BSA) may be done to take up remaining reactive sites.

6. Combine the protein solution with particles and mix thoroughly.

7. React at room temperature 2–4 hours with mixing.

8. Wash beads with coupling buffer and resuspend in the same buffer containing 100 mM

of an amine-containing hydrophilic quenching molecule to block excess reactive sites

(i.e., ethanolamine or Tris).

9. Wash beads and resuspend in an appropriate buffer for storage.

4.4. Coupling to Amine Particles

Primary amine-containing polymeric particles are available from a number of manufacturers

and have either aliphatic or aryl amine groups on their surface. Occasionally, a particle type

may have secondary or tertiary amines present, but these should be avoided for covalent cou-

pling, as primary amines typically give better reaction yields than secondary amines and terti-

ary amines are unreactive.

Primary amine particles may be used for covalent immobilization using a number of reac-

tion routes ( Figure 14.8 ). A carbodiimide-mediated coupling process may be used, as described

above for carboxylate particles, but this time it is done by activation of a carboxylate group

on the ligand and subsequent amide bond formation with the amines on the particles. A single

step EDC reaction strategy will work well for small ligands containing one or more carboxy-

lates with no amines. However, for carbodiimide-mediated protein coupling to amine parti-

cles, the protein ﬁ rst should be activated with EDC and sulfo-NHS according to the method of

Grabarek and Gergely (1990) and then the activated intermediate added to the amine particles

for conjugation (see Chapter 3, Section 1.2 for the activation protocol). This type of two-step

reaction will prevent protein polymerization during the coupling process. Add a quantity of

activated protein to the amine particles to result in a 1–10  molar excess over the total molar

quantity of amines present on all the particles used in the reaction.

4.5. Coupling to Amine Particles Using Crosslinking Agents

Another possible route to coupling ligands to amine particles is to use a bifunctional crosslink-

ing agent to react with the amines and provide another reactive group at the other end to couple

with the ligand. In this approach, virtually any reactive group desired can be formed on the

4. Polymeric Microspheres and Nanospheres 599