Evaluation of different linker regions for multimerization and coupling chemistry for immobilization of a proteinaceous affinity ligand

Martin Linhult, Susanne Gülich, Torbjörn Gräslund, Per-Åke Nygren and Sophia Hober1

Department of Biotechnology, Royal Institute of Technology (KTH), AlbaNova University Center, SE-106 91 Stockholm, Sweden

1 To whom correspondence should be addressed. e-mail: sophia{at}biotech.kth.se


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Alkaline conditions are generally preferred for sanitization of chromatography media by cleaning-in-place (CIP) protocols in industrial biopharmaceutical processes. The use of such rigorous conditions places stringent demands on the stability of ligands intended for use in affinity chromatography. Here, we describe efforts to meet these requirements for a divalent proteinaceous human serum albumin (HSA) binding ligand, denoted ABD*dimer. The ABD*dimer ligand was constructed by genetic head-to-tail linkage of two copies of the ABD* moiety, which is a monovalent and alkali-stabilized variant of one of the serum albumin-binding motifs of streptococcal protein G. Dimerization was performed to investigate whether a higher HSA-binding capacity could be obtained by ligand multimerization. We also investigated the influence on alkaline stability and HSA-binding capacity of three variants (VDANS, VDADS and GGGSG) of the inter-domain linker. Biosensor binding studies showed that divalent ligands coupled using non-directed chemistry demonstrate an increased molar HSA-binding capacity compared with monovalent ligands. In contrast, equal molar binding capacities were observed for both types of ligands when using directed ligand coupling chemistry involving the introduction and recruitment of a unique C-terminal cysteine residue. Significantly higher molar binding capacities were also detected when using the directed coupling chemistry. These results were confirmed in affinity chromatography binding capacity experiments, using resins containing thiol-coupled ligands. Interestingly, column sanitization studies involving exposure to 0.1 M NaOH solution (pH 13) showed that of all the tested constructs, including the monovalent ligand, the divalent ligand construct containing the VDADS linker sequence was the most stable, retaining 95% of its binding capacity after 7 h of alkaline treatment.

Keywords: affinity chromatography/capacity/protein G/purification/serum albumin-binding domain


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Human serum albumin (HSA) has been extensively studied and has found widespread use in both therapeutic and biotechnological applications. At a concentration of about 40 g/l, HSA is the most abundant protein in blood and has several important functions, including a contribution to the colloidal osmotic blood pressure, transportation of fatty acids and binding of therapeutic drugs (Curry et al., 1998Go). HSA is traditionally obtained by fractionation from human plasma. However, the potential hazard of contamination with human viruses has motivated the development of recombinant production systems in yeast (Quirk et al., 1989Go). Purification of a recombinantly produced protein in large-scale industrial processes is generally expensive and time consuming. The overall purification procedure often includes several consecutive unit operations and different chromatographic procedures before the final product is obtained. The introduction of an affinity chromatographic step early in the process for initial product capture and concentration can significantly reduce the number of necessary purification steps (MacLennan, 1995Go). However, such approaches have been hampered primarily owing to the lack of suitable affinity ligands for a large number of products.

We have previously described the use of an albumin-binding domain (ABD) derived from streptococcal protein G for efficient capture of HSA using affinity chromatography. Protein engineering was used to replace alkali-sensitive residues in ABD to create a new ligand, denoted ABD*, that meets industrial requirements for column cleaning-in-place (CIP) procedures based on alkaline exposure (Asplund et al., 2000Go) and therefore has potential for use in large-scale HSA purification (Gülich et al., 2000Go).

The aim of this study was to develop second-generation ligands based on the ABD* protein. Using genetic engineering, head-to-tail dimeric versions have been constructed and further equipped with a unique C-terminal cysteine residue providing a tool for directed coupling of ligands to chromatographic resins. The effect on binding capacity from the dimerization and alternative coupling chemistries has been investigated by both biosensor and affinity chromatography techniques. In addition, the influence on the alkaline stability through the use of three different connective linker sequence candidates has been evaluated.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
DNA constructions

For dimerization of ABD*, the plasmid pTrpABD* (Gülich et al., 2000Go) was used as template in a standard polymerase chain reaction (PCR) protocol. The same vector was used for expression of the monovalent ABD*. Oligonucleotides were purchased from Interactiva (Ulm, Germany) and the cloning was performed using standard protocols according to Sambrook et al. (Sambrook et al., 1989Go). Before dimerization of ABD*, a PCR step to introduce a C-terminal cysteine to allow directed immobilization using the free C-terminal thiol was carried out. The PCR fragment was inserted into plasmid pTrpABD* using the XbaI and PstI restriction sites, to generate pTrpABD*cys. The non-palindromic restriction enzyme AccI was used to steer head-to-tail dimerization of ABD. For the first divalent constructs, ABD*dimerA and ABD*dimerB, the ABD fragments were equipped with AccI sites at both the 3'- and 5'-ends in the PCR step. For the construction of the ABD*dimerB, the asparagine in the pre-existing linker sequence VDANS present at the N-terminus of ABD* was substituted for an aspartatic acid in the PCR step to yield the VDADS linker. This PCR product was ligated into the pGEM-T vector system I (Promega, Madison, WI) and sequenced.

The pGEM-T plasmid carrying the correct ABD* fragment was restricted and ligated in a head-to-tail fashion into plasmids pTrpABD*A and pTrpABD*B predigested with AccI, giving rise to pTrpABD*dimA and pTrpABD*dimB, respectively. Plasmid pTrpABD*dimC was constructed by using two-step PCR (Higuchi et al., 1988Go) with pTrpABD*dimA as template. The gene fragment encoding ABDdimerC was purified and introduced into pTrpABD* cleaved with XbaI and PstI. A MegaBACE 1000 DNA Sequencing System (Amersham Biosciences, Uppsala, Sweden) was used to verify the correct sequence of the inserted fragments. MegaBACE terminator chemistry (Amersham Biosciences) was utilized according to the supplier’s recommendations in a cycle sequencing protocol based on the dideoxy method (Sanger et al., 1977Go). The ligation mixtures were transformed to Escherichia coli strain RR1{Delta}M15 (American Type Culture Collection, Rockville, MD), which was also used for expression of the gene products.

Production and purification

E.coli cells harboring the different ABD* variants were used to inoculate 20 ml of tryptic soy broth (30 g/l) (Difco, Detroit, MI) supplemented with 5 g/l yeast extract (Difco) and 50 mg/l kanamycin monosulfate (LabKemi, Stockholm, Sweden) and were incubated overnight at 37°C. A 5 ml volume of the overnight culture was used to inoculate 500 ml of fresh medium. The culture was incubated overnight at 37°C and the cells were harvested by centrifugation (5000 g for 10 min). After resuspension in TST buffer (25 mM TRIS–HCl, pH 7.5, 150 mM NaCl, 1.25 mM EDTA, 0.05% Tween 20), the cells were disintegrated by sonication (Sonics and Materials, Danbury, CT). For clarification a centrifugation step was carried out (30 000 g for 20 min) and the lysate was filtered (0.45 µm) (Millipore, Bedford, MA).

Proteins were purified using HSA affinity chromatography on an ÄKTA Explorer 10 (Amersham Biosciences) purification system. After washing with TST buffer and 5 mM NH4OAc, pH 5.5, the bound product was eluted with 0.5 M HOAc, pH 2.8. The protein was detected by absorbance measurements at 280 nm and relevant fractions were lyophilized.

The homogeneity was analyzed by SDS–PAGE on high-density gels (Amersham Biosciences) using the Phast system (Amersham Biosciences) under reducing conditions and stained with Coomassie Brilliant Blue (Amersham Biosciences).

Biospecific interaction analysis

A BIAcore 2000 instrument (BIAcore, Uppsala, Sweden) was used for real-time biospecific interaction analysis. Lyophilized protein samples were dissolved in HBS buffer (10 mM HEPES, 0.15 M NaCl, 3.4 mM EDTA, 0.005% surfactant P20, pH 7.4) and filtered (0.45 µm) (Millipore). ABD* and ABD*dimerA were immobilized by N-hydroxysuccinimide and N-ethyl-N'-(3-diethylaminopropyl)carbodiimide chemistry to the carboxylated dextran layer of a CM5 sensor chip (BIAcore) according to the supplier’s recommendations. The immobilization resulted in ~900 RU for each variant. Duplicate samples of 300 nM HSA (Pharmacia, Stockholm, Sweden) in HBS were injected over the surfaces at a flow rate of 10 µl/min.

Prior to immobilization, the cystein-containing ligands were reduced in 10 mM dithiothreitol (DTT) in TST buffer, followed by buffer exchange on Sephadex G-25 columns (Amersham Biosciences) to 50 mM NH4OAc, pH 4.0. The free thiol of the C-terminal cysteine was used for directed coupling of ABD* and ABD*dimerA to the sensor chip. 2-(2-Pyridinyldithio)ethanamine hydrochloride (PDEA) (BIAcore) was injected over an NHS/EDC activated CM5 sensor chip (BIAcore). The immobilization resulted in ~900 RU for ABD* and ABD*dimerA. Duplicate samples of 300 nM HSA (Pharmacia) in HBS were injected at a flow rate of 10 µl/min.

For all analyses, 10 mM HCl was used to regenerate the surfaces. The data were analyzed using the BIA evaluation 3.0.2b software (BIAcore). The signals from a non-immobilized surface (activated and deactivated) were subtracted.

NaOH treatment of the different constructs analyzed by SDS–PAGE analysis

Lyophilized samples were dissolved in 0.5 M NaOH. After incubation for 180 min at room temperature the proteins were precipitated with acetone. The proteins were then dissolved in 1*RED (20 mM TRIS–HCl, pH 8.0, 1 mM EDTA, 2.5% SDS, 5% ß-mercaptoethanol, 0.01% BFB) and analyzed by SDS–PAGE according to manufacturer’s recommendations (Amersham Biosciences).

Affinity chromatography with CIP treatment

ABD* and the different ABD*dimer variants were covalently coupled to an agarose matrix using thioether chemistry (Amersham Biosciences). The different gels were packed in HR5 columns (Amersham Biosciences) for analysis of the stability towards CIP treatment. An ordinary affinity purification protocol was followed on an ÄKTA Explorer 10 (Amersham Biosciences) instrument. HSA (10 mg/ml) (Pharmacia) in TST was loaded at a flow rate of 1 ml/min. After extensive washing with TST, the protein was eluted with 0.5 M HOAc, pH 2.8, supplemented with 100 mM NaCl. NaCl was included to disrupt protein–protein interactions and thereby obtain a narrower peak. After elution the columns were washed with 0.1 M NaOH for 20 min. This procedure was repeated for 21 cycles to follow the decrease in capacity of the columns. The capacities of the columns were determined by measuring the absorbance of the eluted peak. The amount of eluted HSA was also determined by amino acid analysis (BMC, Uppsala, Sweden).


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
We have previously described the HSA-binding affinity ligand ABD*, which corresponds to one of the serum albumin-binding motifs of streptococcal protein G, which has been further engineered to increase its tolerance towards alkaline solutions (Gülich et al., 2000Go). To construct a second-generation ligand, with a potentially higher binding capacity for the HSA target, divalent versions were constructed by genetic head-to-tail fusion of two ABD* moieties. Three divalent constructs, containing different linker sequences connecting the two ABD* domains, were constructed and analyzed (Figure 1B). The first variant (variant A) contained the connective linker sequence VDANS resulting directly from the head-to-tail cloning procedure in the specific vector used. In the second variant (variant B), the asparagine in the linker was substituted for aspartic acid in order to obliterate the alkaline-sensitive dipeptide sequence Asn–Ser (Geiger and Clarke, 1987Go), resulting in a VDADS linker sequence. In addition, a third variant was constructed (variant C) in which the linker sequence included only glycine and serine residues (GGGSG). Linkers composed of these amino acids have been reported to be proteolytically stable, highly flexible and also successful in sterically separating domains (Argos, 1990Go; Alfthan et al., 1995Go; Helfrich et al., 1998Go; Takeda et al., 2001Go). All constructs were equipped with a C-terminal cysteine residue to allow for thiol-based coupling chemistry.



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Fig. 1. (A) An overview of streptococcal protein G (SPG) (Olsson et al., 1987Go). The albumin- and immunoglobulin-binding regions are separated and consist of three highly homologous domains each. ABD* used in this study is an alkali-stabilized variant of one of the albumin-binding domains, ABD 3 (Gülich et al., 2000Go). (B) The three different linker regions analysed in this study. All variants were produced with an N-terminal leader and linker region of 19 amino acids. Also, the different constructs were produced with an additional C-terminal cysteine for directed coupling.

 
The different constructs were expressed as soluble proteins in E.coli and were recovered by HSA affinity chromatography from cell lysates at levels in the range 10–20 mg/ml overnight cultures in shake flasks.

Biospecific interaction analyses

Biosensor binding studies were performed to elucidate any differences in binding capacity between the monovalent original construct ABD* and the divalent variant, ABD*dimerA. In addition, both a directed (thiol coupling) and a non-directed approach (amine coupling) were investigated in parallel to evaluate the importance of the chemistry used for coupling of the ligand to the sensor chip. Approximately equal amounts [measured in resonance units (RU)] of the different ligands were immobilized, thus corresponding to equimolar amounts of ABD* moieties (i.e. number of domains), presented as either monovalent or divalent constructs. The sensorgrams recorded during injection of HSA as analyte over surfaces carrying the respective ligands coupled by non-directed amine coupling chemistry reveal that a divalent ABD* ligand displays a higher dynamic binding capacity than its monovalent counterpart (Figure 2A).




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Fig. 2. Biosensor analysis of interactions between injected HSA and the monovalent ABD* and one divalent variant ligand (ABD*dimerA), respectively. The ABD* variants were attached to the surface by (A) NHS chemistry (non-directed) or (B) thiol chemistry (directed), respectively. The amount of immobilized ABD* domains is similar in all cases (for NHS chemistry, ABD* 970 RU and ABD*dimerA 900 RU; for thiol chemistry, ABD* 905 RU and ABD*dimerA 894 RU). A 300 nM HSA solution was injected over the different surfaces. Mean values from duplicate samples are shown.

 
In contrast, however, when the two different constructs, monovalent and divalent, were coupled by thiol-based chemistry via their C-terminal cysteine residues, the divalent and monovalent ligands showed similar dynamic binding capacities (Figure 2B). It is also noteworthy that the use of a coupling chemistry which resulted in directed immobilization also resulted in a higher overall binding capacity compared with the use of non-directed amine coupling chemistry (see Discussion).

Column affinity chromatography studies

The different divalent ABD* variants, and also the monovalent ABD* domain, were separately coupled to activated Sepharose via their C-terminal cysteines to form directed thioether linkages. The resulting ligand densities of the chromatography matrices were ABD* 3.0 mg/ml, ABD*dimerA 3.4 mg/ml, ABD*dimerB 5.1 mg/ml and ABD*dimerC 2.6 mg/ml (Table I). These affinity media were subsequently used for HSA-binding capacity studies in a standard affinity chromatography protocol involving washing with TST buffer and elution at low pH. The results showed that columns containing divalent ABD* ligands had similar dynamic binding capacities for HSA as calculated per domain, compared with the column containing the monovalent ABD* ligand (Table I). These results corroborated the biosensor studies and indicated that both ABD* moieties in the divalent constructs were equally accessible for interaction with HSA.


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Table I. Dynamic binding capacity of the different ABD* chromatography matrices
 
Incubation of the divalent proteins in 0.5 M NaOH

The divalent ligand constructs with different connective linker sequences were investigated for their stability towards alkaline sanitization procedures by incubation of the purified proteins in 0.5 M NaOH (pH 13.7) for 3 h, followed by analysis by SDS–PAGE. Figure 3 shows that incubation of the divalent ligands at pH 13 resulted in the appearance of varying amounts of lower molecular weight bands, presumed to correspond to degradation products of full-length ligands. For variant C (Figure 3, lane 6), a relatively strong band which co-migrates with the monovalent ABD* reference protein is observed (Figure 3, lane 7). This suggests that the GGGSG linker sequence in this variant was relatively unstable to the treatment and that degradation of some of the material into monovalent ABD* domains occurred. Alkaline treatment of variant A also resulted in the appearance of some degradation products, but with a more even size distribution (Figure 3, lane 4). Interestingly, variant B showed the least degree of degradation compared with the other two variants, suggesting that this variant was the most stable to the alkaline treatment.



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Fig. 3. SDS–PAGE analysis of the different constructs before and after exposure to 0.5 M NaOH for 3 h. Lane 1, ABD*dimerA before incubation; lane 2, ABD*dimerB before incubation; lane 3, ABD*dimerC before incubation; lane 4, ABD*dimerA after alkaline incubation; lane 5, ABD*dimerB after alkaline incubation; lane 6, ABD*dimerC after alkaline incubation; lane 7, purified ABD* (not incubated in NaOH). Small amounts of dimers of the ligands formed by a disulfide-bridge (via their C-terminal cysteine residues) can be seen (lanes 3–7). Molecular weights of the different domains: ABD*, 5.5 kDa; ABD* with N-terminal extension, 7.2 kDa; divalent constructs, 12.1–12.8 kDa.

 
Column sanitization with NaOH

To investigate if the same performance that the ligand variants demonstrated in 0.5 M NaOH was also observed when coupled to the chromatographic resin, a series of multiple-cycle affinity chromatographic experiments under repeated exposure to 0.1 M NaOH were performed.

Each chromatographic cycle consisted of the application of excess HSA to the columns prior to washing, followed by the measurement of the amount of eluted material to determine any decrease in binding capacity of the different resins. A 20 min pulse of 0.1 M NaOH was applied to the columns between each cycle to achieve a total ligand exposure time of 7 h. Interestingly, the results, summarized in Figure 4, showed that the alkaline stability of the ligands ABD*, ABD*dimerA and ABD*dimerC did not vary significantly, suggesting that the two linker variants in these divalent constructs were not affected to a greater extent than the flanking ABD* domains under these conditions. In fact, these ligands retained as much as 85% of the original capacity after 7 h of exposure to 0.1 M NaOH, Interestingly, the ABD*dimerB showed a stability towards high pH exceeding even the monomeric variant and retained as much as 95% of the capacity after 7 h of exposure to 0.1 M NaOH (see Discussion).



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Fig. 4. Influence of alkaline sanitization on the HSA-binding capacity. The monovalent ABD* ligand and the three divalent ABD* ligands were separately investigated in a column chromatography format for stability to repeated alkaline sanitization (0.1 M NaOH). After injection of a saturating amount of HSA, columns were eluted and the binding capacity was determined. Between each purification round, a pulse of 0.1 M NaOH was injected (see Materials and methods for details). The ligands were immobilized using thioether chemistry by means of the C-terminal cysteine. Diamonds, ABD*dimerB; circles, ABD*dimerA; triangles, ABD*dimerC; crosses, ABD*monomer.

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
In this study, the effect of tandem dimerization on a proteinaceous affinity ligand was investigated in terms of binding capacity and alkaline stability using different connective linker sequences and coupling methods. The ABD* ligand investigated here corresponds to one of three homologous domains present in streptococcal protein G (SPG) in which they are separated by connective sequences of 30 amino acids (Olsson et al., 1987Go; Ståhl and Nygren, 1997Go). Native SPG has earlier been shown capable of simultaneous binding to ~2.5 molecules of HSA showing that under certain conditions the three motifs are sufficiently separated to accommodate three molecules of the relative large (66 kDa) interaction partner (Falkenberg et al., 1992Go). Here, considerably shorter connective linkers were used to construct the different divalent constructs, which potentially could lead to a lower molar binding capacity for steric reasons. Three variants of dimers with different linkers (VDANS, VDADS and GGGSG) were successfully produced without detection of any degradation products. Our results, both from the BIAcore analyses and also from the column experiments, indicate that the function of both domains within the ligand is retained when shortening the linker from 30 to five amino acids. When directed coupling chemistry is used, the ABD* domains bind (on a molar basis) HSA equally well regardless of whether used as monovalent domains or tandemly arranged divalent constructs. Interestingly, recent results on the three-dimensional structure of ABD indicate that one amino acid of the N-terminal linker region could be participating in the first {alpha}-helix of ABD (Johansson et al., 2002Go). However, three of the C-terminal amino acids in the domain are suggested to be flexible. Therefore, the flexible part between the two domains is probably seven amino acids, which is similar in length to the connecting regions in protein A (Uhlen et al., 1984Go). However, a significant effect from dimerization of the ABD* was seen when ligands were immobilized to a sensor surface using amine coupling chemistry. Compared with a monovalent ABD* ligand, the divalent counterpart showed an improved binding capacity for HSA (on a domain basis). It could be expected that the coupling chemistry used could result in a loss of binding activity for both types of ligands due to recruitment for attachment of residues involved in binding or by a non-favorable orientation of the ligand. However, to immobilize the same number of ABD* domains when presented in a divalent format, fewer attachment points are needed, resulting in a higher fraction of domains with retained binding activity. This is also supported by the results obtained when the two different ligands were immobilized to the sensor surface using coupling chemistry directed to the C-terminal end, where both ligands showed similar HSA-binding characteristics.

The three divalent constructions with different linker sequences (VDANS, VDADS, GGGSG) were evaluated for use in affinity chromatography. Both alkaline incubation and column chromatographic/alkaline sanitization experiments indicated that all three ligands possess a remarkably overall high stability, but that the ligand containing the VDADS linker was the most stable under high-pH conditions. Surprisingly, the column chromatographic experiment showed that the divalent ligand containing this linker sequence was even more stable than the monovalent ABD* ligand. The reason for this remains unclear, but it cannot be ruled out that the two domains in a divalent ligand can excert a mutual stabilizing effect related to refolding after exposure to denaturing conditions (Robinson and Sauer, 1998Go; Wenk et al., 1998Go). Hence it is not only the linker length and composition but also the function of the separate domains within the protein that are essential for the stability of the ligand. A recently published analysis of different naturally occurring inter-domain linkers shows that the most frequently occurring length is 6–10 amino acids. However, both length and structure differ owing to the properties of the connected domains (George and Heringa, 2003Go). Hence the choice of linker region, i.e. length and composition, is important when connecting two functional domains. Moreover, the choice of coupling chemistry is also of great importance in order to retain the functionality of the connected domains. Our data show that the use of directed coupling results in twice as many active domains for the divalent constructs compared with non-directed coupling. Notably, the difference between directed and undirected coupling is even greater for the monovalent versions of the linker domains.

These results clearly demonstrate the importance of the choice of coupling chemistry and linker sequence to achieve an effective presentation of the functional domain to its target molecule. In addition, they also demonstrate that the designed ABD*dimer with the VDADS linker is highly tolerant towards alkaline treatment and could therefore be a very suitable ligand candidate for the large-scale purification of HSA.


    Acknowledgements
 
We are very grateful to Gunnar Hagström (Amersham Biosciences) for coupling of the ligands to the chromatographic matrix. Finally, we thank Professor Stefan Ståhl (KTH) for fruitful discussions. This project was supported by Affibody AB and Amersham Biosciences.


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 Introduction
 Materials and methods
 Results
 Discussion
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Received May 12, 2003; revised October 8, 2003; accepted October 21, 2003





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