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Ind. Eng. Chem. Res. 2005, 44, 8554-8559
Derivatization of Azlactone-Functional Supports with Small Ligands. Strategies for Control of Ligand Density Jerald K. Rasmussen,* Raymond M. Gleason, Dean S. Milbrath, and Robin L. Rasmussen 3M Corporate Research Laboratories, 3M, 3M Center, 201-2N-20, St. Paul, Minnesota 55144
The derivatization of azlactone-functional beads with small amine-functional ligands was studied with the objective of controlling coupled ligand density. When weakly acidic or weakly basic ligands were coupled in the presence of quenchers, ligand density was readily and reproducibly controlled by the molar ratio of ligand to quencher in the coupling solution. In addition, protein ion exchange capacities were linearly related to coupled ligand densities as well as to molar ratios, even at densities where multiple ionic interactions with the protein were to be expected. These results attest to the versatility of this coupling chemistry and may have broad application for the preparation of derivatized supports useful for large-scale protein separations. 1. Introduction Derivatization of polymeric supports is central to the preparation of various types of diagnostic, chromatographic, and immobilized enzyme media. Coupling of ligands to supports, i.e., covalent attachment of specific molecules or functional groups, is necessary to impart to those supports the ability to effect the separation, identification, and/or purification of molecules of interest. Techniques for controlling the concentration or density of ligands on a polymeric support depend to a certain extent on whether the ligand is a small molecule or a macromolecule (protein or enzyme), but generally fall into one of four categories: (a) Manipulation of reaction conditions, which “activate” the matrix, i.e., which introduce a reactive group, which can couple to the ligand. This often involves varying the concentration of “activating reagents”, reaction time, reaction temperature, pH, or combinations of these variables.1-4 (b) Manipulation of reaction conditions during coupling of the ligand to the support. This may involve varying the concentration and/or total amount of ligand the support is challenged with,2,5-7 ionic strength, and/or type of salt in the coupling buffer,6,8-10 as well as the variables of time, temperature, pH, etc., mentioned above. (c) Manipulation of the amount of reactive or “activatable” functional group incorporated into the polymer support by varying polymer composition at the time of its formation, i.e., during the polymerization.11-13 (d) Manipulation of the amount of ligand incorporated into the polymer by preparation of a polymerizable ligand monomer and varying the concentration of this monomer in the monomer feed during polymerization.14,15 For the most part, the above techniques for controlling ligand concentration on polymeric supports are quite difficult to apply in a practical and reproducible manner, primarily because of the large number of variables that must be simultaneously controlled. This is especially true of the first two techniques, in which the efficiencies * To whom correspondence should be addressed. Tel: 651736-4946. Fax: 651-737-2590. E-mail:
[email protected].
of the reactions (i.e., extent of desired reaction as opposed to competing side reactions) are strongly influenced by reaction conditions. Technique “c” seems to offer some degree of control, although one must subsequently apply the techniques of “a” and/or “b” in a second step to couple the ligand. Technique “d” would appear to provide exact control of ligand density until one realizes that many of the ligands which are useful for diagnostic applications and chromatographic separations contain functional groups which are incompatible with conditions necessary for formation of the desired polymer (e.g., they are unstable under the contemplated polymerization conditions, or they interfere with the polymerization reaction, such as by inhibition of polymerization). With regard to the chromatography of biomacromolecules, the above studies, and others,16-19 have suggested that ligand density and/or ligand distribution may profoundly affect capacity, retention, resolution, and selectivity of the support. In the more specific case of immunoaffinity chromatography, in which the ligand is a high molecular weight protein (an antibody), low antigen binding efficiencies often observed upon immobilization have been attributed to the concerted actions of surface density of antibody, multipoint attachment of antibody to the support, undesirably restrictive conformations imposed by covalent attachment, steric effects, and orientation effects.20,21 Cross-linked, hydrophilic, porous supports bearing azlactone functional groups have been shown to exhibit excellent capacities for the covalent immobilization of protein ligands (Scheme 1).6,11 Those studies also described some of the effects observed on the density of coupled ligand when varying reaction conditions such as pH, ionic strength, coupling time, ligand concentration, and azlactone content of the support. Subsequently, work in our group centered around improving the quality of immobilization of protein ligands.22,23 The goal of this work was to overcome the commonly encountered problem in affinity chromatography of decreasing specific activity of the ligand for its ligate with increasing ligand density.2,7,19,21 Coupling of protein A in the presence of a polyanionic salt and an azlactone quencher was shown to lead to affinity supports having higher molar ratios for IgG bound/protein
10.1021/ie0402867 CCC: $30.25 © 2005 American Chemical Society Published on Web 04/22/2005
Ind. Eng. Chem. Res., Vol. 44, No. 23, 2005 8555 Scheme 1
A coupled as compared to supports prepared in the absence of quencher.22 The presence of the quencher, a molecule which competes with the protein ligand for the azlactone coupling sites, may influence specific activity of the final support in at least two ways. First of all, it may simply help by ensuring a more even spatial distribution, thus preventing steric hindrance due to proximity of adjacent ligands. Second, it may reduce the number of multiple-site attachments of the ligand to the support, thus reducing the possibility for inactive conformations. Additional studies have shown that even distributions of coupled ligands may be achieved using a two-step approach in the coupling process.23 In the first step, the ligand is allowed to permeate the porous support under conditions where the rate of the coupling reaction is very slow. In a second step, reaction conditions are changed to cause rapid coupling to the support. Affinity media prepared by this technique have dramatically improved specific activities and maintain those specific activities to much higher ligand densities than previously observed. These results are again explainable in terms of reducing steric hindrance. The present study was undertaken to develop a simple method for the control of the density of small, low molecular weight ligands on azlactone supports. By conducting ligand immobilization in the presence of an azlactone quencher, a linear relationship is observed between coupled ligand density and the molar or equivalent ratio of ligand to quencher used in the coupling reaction. In addition, similar relationships are observed between chromatographic performance and ligand density or molar ratio. These relationships, to our knowledge, have not been described previously with other supports. 2. Experimental Section 2.1. Materials. Derivatized supports were prepared, unless otherwise noted, using Emphaze Biosupport Medium AB 1 (azlactone-functional bead having an active functionality of approximately 40-45 µmol/mL of swollen support, available from 3M, St. Paul, MN, and distributed under the UltraLink tradename by Pierce Biotechnology, Inc., Rockford, IL).24 Ligand, quencher, and buffer solutions were all prepared, unless otherwise noted, in deionized water. When necessary, pH was adjusted with either 10 M sodium hydroxide or 12 M hydrochloric acid, as appropriate. All chemicals, buffer salts, etc., were commercial materials purchased from Aldrich Chemical Co., Milwaukee, WI, or Sigma Chemical Co., St. Louis, MO, and used as received. 2.2. Procedures. Cation Exchange Capacity. A 0.8 × 4 cm polypropylene disposable chromatography column (Poly-Prep Column, Bio-Rad Laboratories, Hercules, CA) was packed with 1 mL of derivatized bead support. The column bed was equilibrated by washing with 10 mL of loading buffer, 0.01 M MOPS (4-mor-
pholinepropanesulfonic acid)/pH 7.5, and then loaded with 10 mL of protein solution (chicken egg white lysozyme, approximately 95% purity, Sigma Chemical Co., 12 mg/mL in the MOPS buffer), collecting the flow through fraction. Unbound lysozyme was washed off with 30 mL of the MOPS buffer (three 10 mL fractions). Finally, bound protein was eluted with 15 mL of 1 M NaCl in MOPS buffer. Protein recovered in the various fractions was determined by measuring the UV absorbance at 280 nm using a Hewlett-Packard Diode Array Spectrophotometer, Model 8452A, and compared to a standard curve prepared using pure lysozyme. The amount of protein recovered in the NaCl eluate was equated to the cation exchange capacity for the support. Anion Exchange Capacity. The procedure used was identical to that above for cation exchange capacity except that the protein loaded was bovine serum albumin (BSA, fraction V, 96-99% purity, Sigma Chemical Co., 12 mg/mL in MOPS buffer). Pure BSA (Albumin Standard, Pierce Chemical Co., Rockford, IL) was used to construct the standard curve. Amine Functionality Determination. The quantity of immobilized amine ligand was analyzed as follows: A sample of derivatized beads (8-10 mL) was washed successively (using a fritted glass funnel, filter flask, and water aspirator setup) with 150 mL of deionized water, 150 mL of 0.1 M HCl, and 150 mL of 0.0001 M HCl (3-50 mL portions each). Following the final wash, vacuum was maintained until the majority of the liquid had been removed and only a damp filter cake remained. The filter funnel with filter cake was attached to a clean 250 mL filter flask, and the ionically bound chloride ions were displaced by washing with two 20 mL portions of 10% (w/w) sodium sulfate. During each washing step, the beads were suspended and allowed to stand in the solution for 1 min prior to applying vacuum. The combined sodium sulfate filtrates were mixed with 1 mL of 5% (w/w) potassium chromate, stirred vigorously with a magnetic stirrer, and titrated with 0.1000 M silver nitrate to the faint red end point, noting the volume of titrant required. A sample of underivatized beads was used to determine a blank titration volume. The difference between the sample volume and the blank volume was used to calculate the amine content in µmol/mL of bead support. 2.3. Support Derivatizations. Preparation of Carboxyl-Functional Beads. The following solutions were prepared:
ligand solution - 1.0 M aspartic acid, pH 9.0 quencher solution - 3.0 M ethanolamine, pH 9.0 Mixtures of the two solutions were then prepared to provide coupling solutions containing various molar ratios of aspartic acid to ethanolamine; 5 mL of each mixture were added to 125 mg of Emphaze beads and vortexed, and the resultant slurry was allowed to react at room temperature with end-over-end agitation for 2 h, filtered, and washed with deionized water (3 × 10 mL), 0.1 M HCl (10 mL), and finally deionized water until the filtrate was of neutral pH. The results of the derivatizations were assayed by measuring cation exchange capacity for lysozyme. Preparation of Amine-Functional Beads from 1,6-Hexanediamine. The following solutions were prepared:
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Ind. Eng. Chem. Res., Vol. 44, No. 23, 2005
ligand solution 0.5 M 1,6-hexanediamine, pH 11.0 quencher solution - 1.0 M ammonia, pH 11.0 Mixtures of the solutions were prepared (20 mL each), reacted with Emphaze beads (1.00 g, corresponding to 8 mL when hydrated) for 2 h at room temperature, and then worked up by washing with deionized water until the filtrate was of neutral pH. The quantity of immobilized amine ligand and the anion exchange capacity were determined as described above. Preparation of DEAE Beads. DEAE supports were prepared from Emphaze beads (1.25 g, corresponding to 10 mL when hydrated) by procedures similar to the above using the following solutions (prepared without adjustment of pH):
ligand solution - 1.0 M 2-diethylaminoethylamine quencher solution 1.0 M ethanolamine or 1.0 M ammonia Preparation of Amine-Functional Beads from Ethylenediamine. Derivatized supports were prepared as described for DEAE supports except that the ligand solution was 0.5 M ethylenediamine. Preparation of an HIC Support. The following solutions were prepared:
ligand solution - 0.5 M benzylamine, pH 11.0 quencher solution - 0.5 M ethanolamine, pH 11.0 Mixtures of solutions were prepared and reacted with Emphaze beads as in previous examples. The samples were evaluated using a Waters Delta Prep 3000 chromatograph equipped with a Waters Lambda Max UV spectrophotometer and Maxima data acquisition software. Bead samples were packed in a Waters AP-1 column to a bed height of 1.3 cm (1.0 mL total volume). The binding buffer consisted of 1.5 M ammonium sulfate and 0.05 M sodium phosphate, pH 7.1; the elution buffer consisted of 0.05 M sodium phosphate, pH 7.1; the sample protein solution consisted of chicken egg white lysozyme dissolved at a concentration of 5 mg/mL in the binding buffer. Five milliliters of sample protein solution was loaded onto the column at a rate of 1 mL/min. The column was then washed for 5 min with binding buffer at a rate of 1 mL/min and then for 10 min at 2 mL/min. The buffer was then changed to the elution buffer in a single step. Elution of the bound protein was monitored spectrophotometrically (280 nm) until the optical density of the eluted fractions returned to baseline. A sample derivatized with 20 mL of ligand solution was found to bind almost all of the applied protein, and the elution removed the bound protein in a sharp, symmetrical peak with a maximum optical density of about 0.7. A second sample, derivatized with a 50:50 mixture of ligand and quencher solutions, showed a large amount of unbound protein in the flow through fractions, while the elution peak for the bound protein showed a maximum optical density of about 0.35. Preparation and Derivatization of OxiraneFunctional Beads. Oxirane beads were prepared as follows: A 1-L creased, round-bottomed flask equipped with a mechanical stirrer (stirring rate 450 rpm), nitrogen gas inlet, thermometer, and condenser was charged with toluene (188 mL), 0.133 g of poly(isooctylacrylate-
Figure 1. Cation exchange capacity of aspartic acid/ethanolamine supports as a function of mole fraction of aspartic acid in the coupling solution.
co-acryloylaminoisobutyramide), heptane (348 mL), and glycidyl methacrylate (0.72 mL). This mixture was stirred and heated to 35 °C while sparging with nitrogen. To the stirring mixture was added a solution of methylenebisacrylamide (13.3 g), 2-propanol (90 mL), sodium persulfate (0.55 g), and deionized water (60 mL). After stirring for an additional 5 min, tetramethylethylenediamine (0.55 mL) was added to initiate polymerization. Polymerization was allowed to continue for a total of 4 h, and then the resultant beads were filtered, washed with acetone three times, and dried under vacuum overnight to produce oxirane beads having approximately 40 µmol of epoxide functionality per milliliter of support. Beads from a 38-106 µm sieve cut were used in the derivatization experiments. 1.0 M solutions of 2-diethylaminoethylamine and ethanolamine were used to derivatize 1.0 g samples of the above oxirane beads. Evaluation was conducted as in previous examples. 3. Results and Discussion Coupling of small molecules to support materials has been used to provide a variety of ion exchange, hydrophobic interaction, reverse phase, chiral, and affinity chromatography supports as well as functionalized materials for other applications.25 Reaction conditions leading to less than the maximum amount of coupling to azlactone-functional supports could be used, as has been done previously with other coupling chemistries,1-15 to provide supports with varied ligand densities. The problems associated with this approach as mentioned above, and especially potential hydrolysis of residual azlactones to carboxylic acid groups, led us to search for a better method for ligand density control, a method with potential for reproducible scale-up and manufacturing. Our ability to couple proteins in the presence of quenchers22 and a knowledge of conditions leading to efficient coupling of low molecular weight amines26 suggested the possibility of using mixtures of ligand and quencher to control the concentration of ligand coupled to azlactone-functional beads. In an initial study, we derivatized Emphaze AB 1 beads by reaction with mixtures of aspartic acid (a potential weak cation exchange ligand) and ethanolamine (a hydrophilic quencher) at pH 9.0. When direct assay of levels of ligand incorporation by titration of the carboxyls led to inconsistent results, we turned to an indirect assay, cation exchange capacity for lysozyme. The results are shown in Figure 1. Regression analysis indicated that ion exchange capacity correlated to molar ratio of ligand to quencher in the coupling solution with an r2 ) 0.977.
Ind. Eng. Chem. Res., Vol. 44, No. 23, 2005 8557 Table 1. Preparation of Amine-Functional Beadsa ligand
quencher
ligand (equivalent fraction)
ligand density (µmol/mL)
BSA capacity (mg/mL)
0.05 0.1 0.2 0.4 0.5 0.6 0.8 1.0
5.2 12.3 18.8 25.6 29.6 29.0 34.4 41.5
0.0 10.5 11.7 20.8 24.4 25.4 31.2 35.2
16.4 15.2 21.1 28.4 30.2 32.7 26.8 44.1
0.3 0.6 2.8 17.5 15.4 15.7 9.2 25.8
14.1 27.2 23.7 39.1 42.4
0.7 7.7 10.2 18.4 19.0
1,6-hexanediamine
ammonia
2-diethylaminoethylamine
ethanolamine
[3.0 M]
none none ammonia none
0.1 0.2 0.5 0.8 1.0 1.0 0.5 1.0
ammonia ethanolamine ammonia none none
0.2 0.5 0.5 1.0 1.0
none
1,2-ethylenediamine
[1.0 M] a
Except as noted, concentrations of ligand solutions were as noted in the Experimental Section; mixtures of ligand and quencher were prepared such that the final primary amine concentration in the coupling solution was 1.0 M.
tions which indicated that, above a ligand density of about 70 µmol/g, ligands were in close enough proximity that multiple interactions with the protein molecule were possible (it was estimated that at 72 µmol/g the distance between the two nearest ligands was 15 Å); thus, increasing ligand density above this level had little effect on protein binding capacity. Data from an evaluation of commercially available cation exchange supports27 seems to be in agreement with Wu and Walters’ conclusions. Following Wu and Walters’ approach, we calculated the average distance between ligands using the equation Figure 2. Anion exchange capacity of 1,6-hexanediamine derivatized supports as a function of amine ligand density.
S ) (A/CN)1/2
Encouraged by these results but unsure of their implications because of the lack of a direct assay for ligand incorporation, we next investigated the preparation of amine-functional beads (potential weak anion exchange supports). Emphaze beads were derivatized with mixtures of 1,6-hexanediamine (HD) and ammonia (Table 1). In this case, a more direct assay of coupled ligand density was possible by conversion to the amine hydrochloride, displacement of the chloride ion with sulfate, and titration of the displaced chloride with silver nitrate. Multiple analyses (six identical derivatizations) established reproducibility, with standard deviations of 1.9 µmol/mL for the amine content measurement and 1.2 mg/mL for the BSA capacity measurement. Once again linear relationships were observed, this time between ion exchange capacity and ligand density (r2 ) 0.979, Figure 2) as well as between ion exchange capacity and molar ratio (r2 ) 0.932). These results are in marked contrast to observations made by Wu and Walters4 upon conversion of diolbonded silica to cation exchange matrixes through reaction with diglycolic anhydride. These workers found a linear relationship between capacity factor (k′) for a small molecule, benzylamine, and ligand density in the low to intermediate ligand density region (up to about 150 µmol/g) but a nonlinear relationship between lysozyme binding capacity and ligand density in the same region. The results were explained using calcula-
where S is the average distance between two nearest ligands, A is the surface area of the support (ca. 350 m2/g for Emphaze beads24), C is the ligand density (corrected for the swell volume of 8 mL/g24), and N is Avogadro’s number. This allowed estimation of the range 13.3-37.4 Å average distance between ligands for the HD resins shown in Figure 2. Although this is certainly well within the range of multisite binding to BSA (BSA has been found28 to resemble a prolate ellipsoid in solution having major and minor axes of 140 and 40 Å, respectively), we see a significant increase in capacity with increase in ligand density. From the slope of the curve, we can estimate approximately 1.0 mg of capacity/µmol of ligand added, nearly 10-fold that estimated from Wu and Walters’ data. Thus, it appears that ligand sites on the Emphaze matrix may be more efficiently utilized for protein binding than on some other supports. Since Wu and Walters recognized that heterogeneity of binding sites probably contributed to some of their observed results, a more uniform distribution of ligand sites might contribute in our case. It would be of interest to extend our studies to much higher ligand densities to see whether the observed relationships would continue to hold. That study, however, must await the preparation of supports with higher azlactone content. The next step was to extend the results to the preparation of a more conventional weak anion-ex-
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Ind. Eng. Chem. Res., Vol. 44, No. 23, 2005 Table 2. Derivatization of Oxirane Beads with Diethylaminoethylaminea DEAE (mole fraction)
ligand density (µmol/mL)
BSA capacity (mg/mL)
0.2 0.5 1
32.3 33.6 39.1
22.9 25.4 26.7
a Derivatizations were conducted with mixtures of 2-diethylaminoethylamine (DEAE) and ethanolamine at total primary amine concentrations of 1.0 M in deionized water.
Figure 3. Anion exchange capacity as a function of ligand density for 2-diethylaminoethylamine (0) and 1,2-ethylenediamine (2) derivatized beads.
change resin. Azlactone beads were reacted with 2-diethylaminoethylamine in an admixture with varying amounts of ethanolamine, leading to DEAE-functional supports (Table 1). Analysis of the exchange capacities for bovine serum albumin (BSA) revealed a linear relationship having approximately the same slope as that observed for the HD beads; however, the line was displaced nearly 10 mg/mL lower in exchange capacity at equivalent ligand densities. Several factors were considered as possible explanations for this effect: (1) the difference in amine functionality (primary vs tertiary) in the two series; (2) the difference in quenching agent (ammonia vs ethanolamine) which, in turn, might affect the hydrophilic nature of the background support; (3) a spacer effect, which places the ionic group at differing distances (12 vs 8 atoms) from the polymer backbone; and (4) a hydrophobic effect due to the hexamethylene group in the HD series. Subsequent experiments with 2-diethylaminoethylamine/ammonia or 1,2-ethylenediamine (ED)/ammonia and ED/ethanolamine (Table 1 and Figure 3) indicated that all ethylenediamine derivatives obeyed the same relationship regardless of the nature of the amine functionality or of the nature of the quenching agent. A single experiment in which beads were derivatized with 3,6-dioxa-1,8-octanediamine produced a support with a ligand density of 34.1 µmol/mL and an ion exchange capacity of 24.7 mg/mL, seeming to discount the importance of a spacer effect. On the other hand, beads derivatized with 1,10-diaminodecane produced a support with a ligand density of 48.9 µmol/mL and an exchange capacity of 46.9 mg/mL, values which fall right in line with those of the HD series. Although the concentration of primary amine in all of these experiments has been chosen to be at least 20-fold that of the azlactone functional groups, the possibility remains that both ends of a diprimary amine could react with neighboring azlactones. This possibility, however, cannot explain the observations since it is anticipated that such intramolecular reaction would be more prevalent with the longer chain diaminohexane and diaminodecane molecules. One would expect lower ligand densities as a result under comparable reaction conditions with these species as compared to the ED series, yet the opposite was found. Further work may be necessary to develop a complete understanding of these results. Although an extensive study was not undertaken, it appears that our results can be extended to the preparation of hydrophobic interaction supports as well.
Derivatization of beads with a 1:1 molar ratio of benzylamine and ethanolamine produced a support that bound approximately 50% of the amount of lysozyme bound by a support derivatized with only benzylamine when assayed under typical HIC conditions. Finally, we explored the possibility of using our method of controlling ligand density with another coupling chemistry. For this work, we prepared oxiranefunctional beads with approximately the same level of functionality as the azlactone-functional beads. These were then derivatized with mixtures of 2-diethylaminoethylamine and ethanolamine. The results, shown in Table 2, indicated that not only were we unable to control ligand density over a very wide range but also that there was no linear relationship between ligand density and anion exchange capacity for BSA. Although additional experimentation will be necessary to gain a complete understanding, there are several possible explanations for these results: (1) the incorporation of the oxirane monomer may result in a different morphology in the bead structure, and subsequently the oxirane functional group may not be as accessible as is the case with the azlactone; (2) the secondary amine formed upon reaction with an oxirane group is potentially reactive toward a second oxirane functional group; and (3) competitive hydrolysis under the reaction conditions may be more prevalent with the oxirane coupling chemistry. 4. Conclusions Azlactone coupling chemistry appears to offer exceptional control over ligand density and distribution. This appears to be due in large part to a facile coupling reaction with amine nucleophiles and a correspondingly slow competing hydrolysis reaction in aqueous media. This work has shown that with small, amine-functional ligands, ligand density is readily and reproducibly controlled by immobilization in the presence of a quencher. While only a small subset of potential ligands has been examined, it seems reasonable to expect that the results can be expanded to other ligands. With larger protein ligands, which display a much lower intraparticle diffusion rate than do small molecules, additional coupling condition variables such as pH, temperature, and ionic strength provide options for controlling distribution and density of ligands.23 This control over ligand density and distribution should provide one more means for the optimization of chromatographic performance and the improvement of process efficiency. Literature Cited (1) Landgrebe, M. E.; Wu, D.; Walters, R. R. Preparation of chromatographic supports of variable ligand density. Anal. Chem. 1986, 58, 1607.
Ind. Eng. Chem. Res., Vol. 44, No. 23, 2005 8559 (2) Eveleigh, J. W.; Levy, D. E. Immunological Characteristics and Preparative Application of Agarose-based Immunosorbents. J. Solid-Phase Biochem. 1977, 2, 45. (3) Lewis, L. A.; Kip, K. F. Preparation of Ligand-Polymer Conjugate Having a Controlled Number of Introduced Ligands. U.S. Patent 4,968,742, 1990. (4) Wu, D.; Walters, R. R. Effects of stationary phase ligand density on high-performance ion-exchange chromatography of proteins. J. Chromatogr. 1992, 598, 7. (5) Anderson, D. J.; Walters, R. R. Affinity chromatographic examination of a retention model for macromolecules. J. Chromatogr. 1985, 331, 1. (6) Coleman, P. L.; Walker, M. M.; Milbrath, D. S.; Stauffer, D. M.; Rasmussen, J. K.; Krepski, L. R.; Heilmann, S. M. Immobilization of Protein A at high density on azlactone-functional polymeric beads and their use in affinity chromatography. J. Chromatogr. 1990, 512, 345. (7) Fowell, S. L.; Chase, H. A. Variation of immunosorbent performance with the amount of immobilized antibody. J. Biotechnol. 1986, 4, 1. (8) Smalla, K.; Turkova, J.; Coupek, J.; Hermann, P. Influence of Salts on the Covalent Immobilization of Proteins to Modified Copolymers of 2-HydroxyethylMethacrylate with Ethylene Dimethacrylate. Biotechnol. Appl. Biochem. 1988, 10, 21. (9) Wheatley, J. B. Effect of antigen size on optimal ligand density of immobilized antibodies for a high-performance liquid chromatographic support. J. Chromatogr. 1991, 548, 243. (10) Wheatley, J. B.; Schmidt, D. E., Jr. Salt-induced immobilization of proteins on a high-performance liquid chromatographic epoxide affinity support. J. Chromatogr. 1993, 644, 11. (11) Rasmussen, J. K.; Heilmann, S. M.; Krepski, L. R.; Jensen, K. M.; Mickelson, J.; Johnson, K.; Coleman, P. L.; Milbrath, D. S.; Walker, M. M. Crosslinked, Hydrophilic, Azlactone-functional Polymeric Beads: A Two-step Approach. React. Polym. 1991/1992, 16, 199. (12) Hradil, J.; Svec, F. Synthesis of Methacrylate Copolymers with Tributylammonium Groups. Polym. Bull. 1985, 14, 265. (13) Arshady, R.; Ledwith, A. Suspension polymerization and its application to the preparation of polymer supports. React. Polym. 1983, 1, 159. (14) Nowinski, R. C.; Hoffman, A. S.; Houghton, R. L.; Priest, J. H.; Monji, N. Synthesis and use of polymers containing integral binding-pair members. U.S. Patent 4,752,638, 1988. (15) Lee, R. T.; Cascio, S.; Lee, Y. C. A simple method for the preparation of polyacrylamide gels containing thioglycoside ligands. Anal. Biochem. 1979, 95, 260.
(16) Fausnaugh, J. L.; Kennedy, L. A.; Regnier, F. E. Comparison of Hydrophobic-interaction and Reversed-phase Chromatography of Proteins. J. Chromatogr. 1984, 317, 141. (17) Narayanan, S. R.; Crane, L. J. Affinity chromatography supports: a look at performance requirements. Trends Biotechnol. 1990, 8, 12. (18) Hage, D. S.; Walters, R. R.; Hethcote, H. W. Split-peak affinity chromatographic studies of the immobilization-dependent adsorption kinetics of Protein A. Anal. Biochem. 1986, 58, 274. (19) Hearn, M. T. W.; Davis, J. R. Evaluation of factors which affect column performance with immobilized monoclonal antibodies: Model studies with a lysozymesantilysozyme system. J. Chromatogr. 1990, 512, 23. (20) Velander, W. H.; Subramanian, A.; Madurawe, R. D.; Orthner, C. L. The use of Fab-masking antigens to enhance the activity of immobilized antibodies. Biotechnol. Bioeng. 1992, 39, 1013. (21) Spitznagel, T. M.; Clark, D. S. Surface density and orientation effects on immobilized antibodies and antibody fragments. Bio/Technology 1993, 11, 825. (22) Coleman, P. L.; Milbrath, D. S.; Walker, M. M. Biomolecules covalently immobilized with a high bound specific biological activity and method of preparing same. U.S. Patent 5,200,471, 1993. (23) Subramanian, A.; Van Cott, K. E.; Milbrath, D. S.; Velander,W. H. Role of local antibody density effects on immunosorbent efficiency. J. Chromatogr. A 1994, 672, 11. (24) Johnson, P. R.; Stern, N. J.; Eitzman, P. D.; Rasmussen, J. K.; Milbrath, D. S.; Gleason, R. M.; Hogancamp, R. E. Reproducibility of physical characteristics, protein immobilization and chromatographic performance of 3M Emphaze Biosupport Medium AB 1. J. Chromatogr. 1994, 667, 1. (25) Akelah, A.; Moet, A. Functionalized Polymers and Their Application; Chapman and Hall: New York, 1990. (26) Derivatization of 3M Emphaze Biosupport Medium AB 1 With Small Amine Ligands. Applications Note, 3M Bioapplications, 1993. (27) DePhillips, P.; Lenhoff, A. M. Determinants of protein retention characteristics on cation-exchange adsorbents. J. Chromatogr. A 2001, 933, 57. (28) Peters, T. Serum albumin. Adv. Clin. Chem. 1970, 13, 37.
Received for review November 29, 2004 Revised manuscript received March 15, 2005 Accepted March 28, 2005 IE0402867
