Emerging biomaterials for downstream manufacturing of therapeutic proteins

Acta Biomaterialia 95 (2019) 73–90

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Acta Biomaterialia journal homepage: www.elsevier.com/locate/actabiomat

Review article

Emerging biomaterials for downstream manufacturing of therapeutic proteins q Yi Li a, David Stern a, Lye Lin Lock b, Jason Mills b, Shih-Hao Ou a, Marina Morrow a, Xuankuo Xu b,⇑, Sanchayita Ghose b, Zheng Jian Li b, Honggang Cui a,c,⇑ a Department of Chemical and Biomolecular Engineering, and Institute for NanoBioTechnology, The Johns Hopkins University, 3400 North Charles Street, Baltimore, MD 21218, United States b Biologics Process Development, Global Product Development and Supply, Bristol-Myers Squibb, Devens, MA 01434, United States c Department of Oncology and Sidney Kimmel Comprehensive Cancer Center, Johns Hopkins University School of Medicine, Baltimore, MD 21205, United States

a r t i c l e

i n f o

Article history: Received 5 November 2018 Received in revised form 26 February 2019 Accepted 6 March 2019 Available online 9 March 2019 Keywords: Biomaterials Therapeutic proteins Downstream processing Chromatography Affinity precipitation

a b s t r a c t Downstream processing is considered one of the most challenging phases of industrial manufacturing of therapeutic proteins, accounting for a large portion of the total production costs. The growing demand for therapeutic proteins in the biopharmaceutical market in addition to a significant rise in upstream titers have placed an increasing burden on the downstream purification process, which is often limited by high cost and insufficient capacities. To achieve efficient production and reduced costs, a variety of biomaterials have been exploited to improve the current techniques and also to develop superior alternatives. In this work, we discuss the significance of utilizing traditional biomaterials in downstream processing and review the recent progress in the development of new biomaterials for use in protein separation and purification. Several representative methods will be highlighted and discussed in detail, including affinity chromatography, non-affinity chromatography, membrane separations, magnetic separations, and precipitation/phase separations. Statement of Significance Nowadays, downstream processing of therapeutic proteins is facing great challenges created by the rapid increase of the market size and upstream titers, starving for significant improvements or innovations in current downstream unit operations. Biomaterials have been widely used in downstream manufacturing of proteins and efforts have been continuously devoted to developing more advanced biomaterials for the implementation of more efficient and economical purification methods. This review covers recent advances in the development and application of biomaterials specifically exploited for various chromatographic and non-chromatographic techniques, highlighting several promising alternative strategies. Ó 2019 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

Contents 1. 2.

3.

q

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Affinity chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Affinity ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Fusion tags. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Non-affinity chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Ion exchange modalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

74 75 75 77 78 78

Part of the Cell and Tissue Biofabrication Special Issue, edited by Professors Guohao Dai and Kaiming Ye.

⇑ Corresponding authors at: Biologics Process Development, Global Product Development and Supply, Bristol-Myers Squibb, Devens, Massachusetts 01434, United States (X. Xu). Department of Chemical and Biomolecular Engineering, and Institute for NanoBioTechnology, The Johns Hopkins University, 3400 North Charles Street, Baltimore, MD 21218, United States (H. Cui). E-mail addresses: [email protected] (X. Xu), [email protected] (H. Cui). https://doi.org/10.1016/j.actbio.2019.03.015 1742-7061/Ó 2019 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

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4. 5. 6.

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3.1.1. Zwitterionic cellulose beads. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2. Electrospun cellulose nanofibers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Hydrophobic/hydrophilic interaction modalities. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Mixed-mode ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Membrane separation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Magnetic separation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Precipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. Non-affinity precipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1.1. PEG-induced precipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1.2. Polyelectrolytes-induced precipitation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1.3. Self-precipitation with fusion tags . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. Affinity precipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.1. ELP-mediated precipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.2. Peptide-based materials and smart polymers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aqueous two-phase systems (ATPs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions and future perspectives. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix A. Supplementary data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Therapeutic proteins have become one of the largest and fastest growing biopharmaceutics in the last two decades and have received increased attention for the treatment of cancer, cardiovascular diseases, infections, genetic disorders, and other diseases [1,2]. The global protein therapeutics market hit $140.1 billion in 2016, and is predicted to reach $217.6 billion by 2023 [3]. Monoclonal antibodies (mAbs) comprise the largest class of therapeutic proteins and were valued at $85.4 billion in 2015 [4]. Other major therapeutic proteins include fusion proteins, insulin, vaccines, enzymes, cytokines, erythropoietin (EPO), interferon, human growth hormone, and blood clotting factors. The rapidly expanding therapeutic proteins market presents a great challenge to industrial production, especially in downstream processing due to its high cost and limited production capacities. Meanwhile, increasing pressure on downstream process operations is due to the significant growth of upstream titers. More than tenfold increase in titers has been seen over the past 30 years with an average of 3 g/L, while several new products are reported to reach more than 10 g/L [5–7]. In contrast, improvement of downstream processing lags substantially behind the increase in titers, and the associated downstream costs increase dramatically as the purification capacity rises up [5]. The emergence of advanced biomaterials has contributed to addressing the downstream bottleneck by introducing high-

79 79 80 80 80 82 83 83 83 84 84 85 85 85 86 86 86 87 87

performance and cost-effective downstream technologies [5,8,9]. Typically, downstream mAb processing involves a sequence of different chromatographic separations, virus inactivation and filtration, and membrane-based separations [5]. After clarifying cell culture harvest, protein A or other affinity-based chromatography is used to capture the target mAb product, followed by a virus inactivation step [5,10]. Then combinations of polishing chromatography steps, including, but not limited to, ion exchange chromatography (IEX) and hydrophobic interaction chromatography (HIC), are allowed to further reduce impurity levels by utilizing the differences in physical properties such as charge, hydrophobicity, and molecule size [5,11]. Finally, a virus filtration step and an ultrafiltration/diafiltration step are performed to ensure viral clearance and to exchange the product into the formulation buffer, respectively [9,10]. Despite the high media cost and limited loading capacity, affinity chromatography remains the most widely used capture method for large-scale purification [8]. Proteins of interest can be separated from impurities through reversible interactions between the target protein and affinity ligands immobilized on chromatography media. For example, protein A chromatography has been widely used in the purification of mAbs and Fc-fusion proteins [12]. Over the past decades, numerous affinity ligands have been developed to improve the binding capacity, capture efficiency, and ligand stability [13]. Furthermore, affinity tags that are genetically grafted onto target proteins have

Fig. 1. Schemes of representative methods discussed in this review for purification of therapeutic proteins.

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also gained widespread attention, as these pre-defined affinity pairs are independent of each particular target protein [14,15]. Advances have also been made in developing more efficient chromatography media such as natural or synthetic polymers to achieve higher column performance [11]. To overcome limitations of chromatographic procedures for high cost and demanding operations, continuous efforts have also been made to develop nonchromatographic methods including membrane separations [16], magnetic separations [17], and precipitation/phase separations [18,19]. These are promising alternatives to chromatographybased approaches, with advantages including cost-effectiveness and operational simplicity in some applications. Among these non-chromatographic methods, polymer/peptide-based biomaterials have been intensively utilized in the design and construction of membranes, magnetic nanoparticles, and precipitants with high capture efficiency and high selectivity. This article provides an overview of the recent advances in the development and optimization of biomaterials for applications in downstream processing of therapeutic proteins (Fig. 1). Approaches currently used in industry and in development will be discussed. 2. Affinity chromatography 2.1. Affinity ligands Affinity chromatography is the predominant unit operation for large-scale bioprocessing and purification of therapeutic proteins [13]. Despite its high binding specificity to the target protein, affinity chromatography is currently still the downstream productivity bottleneck due to its relatively low resin binding capacity, limited column size amenable to biomanufacturing, and high resin cost [20]. Affinity ligands that are immobilized on the chromatography media can bio-recognize and bind to specific proteins through molecular interactions such as hydrophobic and electrostatic interactions, hydrogen bonding, and van der Waals forces [13]. Important features of an affinity ligand include its affinity and specificity to the target, stability in various solution conditions, ease of immobilization, and target-binding retention following attachment to the matrix [21]. A myriad of affinity ligands can be used for protein purification depending on the biophysical properties of the target protein. Biospecific adsorbents, for example, are

ligands of biological origin that are derived from natural sources and can bind to specific targets such as bacterial immunoglobulin-binding proteins, antibodies, lectins, and nucleic acids [13]. The majority of ligands currently being used in affinity chromatography are biospecific ligands. Table 1 shows some of the vast array of affinity ligands that have gained interest over the last several years for applications in the purification of therapeutic proteins; several others exist which have been reviewed elsewhere, and will not be discussed here [8,22,23]. The most frequently used biospecific ligands for mAb and Fcfusion protein purification processes are protein A, protein G, and protein L, which are all derived from bacterial cell walls [13]. Many species of antibodies can bind to these ligands, yet, their affinities vary depending on the antibody subclass [47,48]. However, the harsh environment during elution and other conditions at extreme pH and exposure to organic solvents or detergents can denature coupled biospecific molecules, often resulting in ligand leaching [21]. The stability and binding capacity of recombinant protein A affinity resins are constantly being improved. For example, Toyopearl AF-rProtein A-650F, a recombinant protein A affinity resin derived from an immunoglobulin G (IgG)-binding domain, has been optimized to withstand alkaline solutions during cleaning and sanitization procedures and displays higher binding capacities, up to 50–70 mg/ml, yielding eluted antibodies of about 99% purity [34]. Additional resins with high binding capacities and high alkaline stabilities are found in GE Healthcare’s MabSelect family, including MabSelect Sure, Sure LX, Sure pcc, and PrismA [25–27]. Within this family of resins, MabSelect Sure pcc and MabSelect PrismA have the highest binding capacities for mAbs and bispecific antibodies, ranging from 58 to 74 mg/mL. These MabSelect resins are derived from the B domain of protein A, giving them a high binding affinity for various mAbs and Fc-fusion proteins, such as immunoglobulins [25–27]. Another approach is to use single-domain camelid antibodies as affinity ligands which are advantageous because of their small size, high affinity, stability, and 3D structure that facilitates binding to various biological epitopes present in enzymes, peptides, hormones, and viruses [49,50]. Ligands for camelid heavy-chain variable domains (VHHs) have been used for chromatographic purification of various molecules such as blood factors, antibodies, and adeno-associated viruses [28]. Reinhart et al. used an

Table 1 Commonly used affinity ligands for protein purification in affinity chromatography. Ligand/Family

Affinity Target

Scaffold/Origin

References

Recombinant protein A derived from an IgG-binding domain MabSelect Sure, Sure LX, Sure pcc, PrismA Camelid VHH

IgG, IgM, and Fab fragments

Fc-region (B domain) and Fab-region (D&E) domains of protein A B domain of protein A

[24]

VHH antibody

[28,29]

Z domain, modified from B domain of Staphylococcal protein A (SpA) Z domain, modified from B domain of SpA Z domain, modified from B domain of SpA

[30]

Anti-EGFR Affibody Anti-TNF-a Affibody Anti-idiotypic Affibody Con A 3-aminophenylboronic acid Repebody Ugi and Triazine de novo ligands Short peptide ligands (HWRGWV, HYFKFD, and HFRRHL) Short cyclic peptide ligands (cyclo[LinkM-WFRHY-K]) Cibacron Blue F-G3A Aptamers (e.g. Mapt2.2CS, MaptH1.1CSO, Nonapta5.1)

Fc-region of IgG IgA, IgG, various blood factors, and adeno-associated viruses EGFR receptor TNF-a receptor Human epidermal growth factor receptor 2 affibody molecule Glycoproteins/carbohydrate residues Glycoproteins/carbohydrate residues IL-6, lysozyme, Fc fragment of IgG Glycoproteins, EPO, IgG (Fc and Fab fragments)

[25–27]

[30] [31] [32] [33] [34] [35–38]

Fc region of IgG

Lectin Boronate Variable lymphocyte receptors protein L (A&C domain), protein G Streptococcal protein G (SpG-III domain) Fc-binding domain of protein A

Fc region of IgG

Fc-binding domain of protein A

[41]

Several enzymes and proteins (non-specific) Variety of small molecules, proteins, membrane-bound receptors, cell surface epitopes

Reactive dye and protein, EPO Nucleotide sequences selected from combinatorial libraries (i.e. SELEX)

[42] [43–46]

[39,40]

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Y. Li et al. / Acta Biomaterialia 95 (2019) 73–90

immobilized camelid anti-human VHH ligand to isolate recombinant immunoglobulin A from Chinese Hamster Ovary cell suspensions with high purity and more than 95% recovery in a single chromatographic step [29]. The problem with single-domain camelid antibody ligands is that they may display nonuniformity upon resin immobilization, resulting in reduced capture efficiency of the target. Furthermore, the large size of the antibody ligands restricts their coupling efficiency to the resins in the column, limiting their binding capacity [13,50]. Affibody molecules are a class of small protein domains that have been used for many biotechnological applications including diagnostic and therapeutic applications [30]. A number of affibody molecules such as anti-epidermal growth factor receptor (EGFR) and anti-tumor necrosis factor (TNF)-a affibodies have been used to specifically capture certain biomolecules [30]. Wallberg et al. introduced an anti-idiotypic affibody molecule produced in Escherichia coli (E. coli), purified, and chemically coupled to a resin in an affinity column [31]. Their affibody, serving as an affinity ligand, can selectively bind to human epidermal growth factor 2 (HER2) with high affinity, suitable recovery, and purity for different variants of HER2 affibody molecules. Each HER2 affibody molecule, designed for future radioimaging applications, had different Cterminal peptide extensions and was successfully recovered with high purity in a single step [31]. This purification platform demonstrates its potential to efficiently recover and purify related variants of a target protein that may have modified biochemical properties, thus avoiding the need for specific recovery methods for each variant of target protein. Another biospecific affinity ligand used for capturing a variety of proteins is the Repebody, named for its naturally occurring antibodies composed of repeat modules [34]. Repebodies have been used to purify myeloid differentiation protein-2 and hen egg lysozyme among other biomolecules. Repebodies can also be used to create molecular binders with high affinity and specificity for several different targets by phage display selection [34]. Glycosylation plays a major role on the biological activity and pharmacological behavior of recombinant proteins. Processing therapeutic glycoproteins requires a heterogenous mixture of glycoforms to be isolated and purified. One way to achieve this is to use ligands with high affinity for various glycoforms. For instance, lectins, which are found in plants, animals, and microorganisms demonstrate specificity for certain types of carbohydrate residues [51]. As a result, lectins are commonly used to isolate and identify glycoproteins, glycopeptides, and oligosaccharides; among them, Concanavalin A (Con A) is predominantly used. Du et al. recently explored an immobilization technique using Con A to design reversible affinity ligands adsorbed on a chromatographic support [32]. By avoiding the use of irreversible ligands on chromatographic supports, the expensive support can be recycled for further purification operations with the ligands discarded once they become inactive after several purification cycles [32]. They accomplished this by using a macroporous cellulose monolith as a chromatographic support with covalently immobilized carboxylated groups and using Cu ions to bridge the modified support structure with Con A affinity ligands. Recoveries of glucose oxidase, the target glycoprotein, ranged from 91 to 95%, depending on the flow velocity of the mobile matrix in the column [32]. Boronic acid ligands present a more economical and stable alternative to lectins for separating cis-diol-containing substances such as nucleotides, nucleic acids, and carbohydrates [33]. 3-aminophenylboronic acid is the predominantly used boronic acid ligand which is commonly immobilized on agarose beads. However, this ligand interacts optimally with glycoproteins at pH values beyond 8.5, which may negatively impact the biological activity of the target molecules [33]. As a result, current studies are focusing on alternative

glycoprotein-binding adsorbents through a biomimetic approach [35,52]. Since most biospecific ligands are derived from bacteria, there is a risk of contamination by viruses, DNA, and pyrogens; thus, further purification is essential, which increases the manufacturing and bioprocessing costs. Compared to biospecific ligands, pseudobiospecific ligands, are more robust, economically feasible, structurally simple, less toxic, and very stable with resistance to punitive sterilization environments. However, pseudo-biospecific ligands are not as selective as biological ligands, and require substantial optimization for each protein to achieve high selectivity, thus limiting their acceptability as a superior purification technique in industry [21]. There are many types of synthetic ligands that can be chemically modified including biomimetic peptoidal, peptidic, and dye-based ligands. Biomimetic peptoidal ligands utilize the physicochemical properties of amino acid residues to interact and bind with the target protein. For example, protein A mimetic peptoidal ligand is commonly used for mAb purification because of its specificity towards the Fc fragment of immunoglobulins. Rational design and synthesis of synthetic ligands like biomimetic triazine and Ugi ligands, known as de novo ligands, have already led to successful purification of several proteins at the bench scale [35–38]. Linear and cyclic short peptide ligands ranging from two to nine amino acid residues have recently been shown to have great potential for protein purification [23]. Because of their smaller molecular weight and short half-lives, linear short peptide ligands are less likely to cause an immune response in case of ligand leakage. Additionally, ligands of this nature can more readily elute at milder conditions due to their moderate interactions with target proteins [53]. Yang et al. reported three linear hexapeptides (HWRGWV, HYFKFD, and HFRRHL) which could bind to the Fc-region of all human subclasses of IgGs with high specificity and affinity [39,40]. Short cyclic peptides have also been used as affinity ligands for IgG purification. Cyclic peptides are resistant to proteolysis, and have higher affinity and specificity compared to short linear peptides, making them attractive candidates as affinity ligands [41]. Rao and coworkers developed a strategy of synthesizing a pool of selected pentapeptides directly on a chromatographic support and found that resins based on their cyclo[Link-M-WFRHY-K] peptide ligand have a robust affinity and high selectivity for Fcbinding to human IgG [23,41]. Furthermore, the use of cyclo [Link-M-WFRHY-K] peptide ligand resulted in a high IgG recovery similar to that of protein A resins, while mitigating the need for harsh elution conditions [41]. Dye-based ligands contain reactive dyes that bind to proteins selectively and reversibly [54]. For example, Cibacron-Blue F-G3A dye has been used as affinity ligands for interferon purification using fast protein liquid chromatography (FPLC) [42]. Interferons are a group of glycoproteins known as cytokines that play an important role in immune defense. Dogan et al. used supermacroporous poly(2-hydroxymethyl methacrylate) (poly(HEMA)) cryogels covalently immobilized with Cibacron-Blue (CB) F-G3A, together known as poly(HEMA)/CB, to specifically adsorb recombinant human interferon-a (rHuIFN-a) with high affinity through interactions among functional groups of the dye and rHuIFN-a [42]. The interferon samples were purified using FPLC which yielded 97.6% purity and 84.6% recovery [42]. Despite the robustness of this platform, there have been concerns regarding the purity, selectivity, leakage, and toxicity of some commercially available dyes which have limited their use in industrial purification systems. Nucleic acid aptamers have also been used as ligands in affinity chromatography for the purification of a wide range of target proteins [45,46]. Aptamers are single stranded DNAs, RNAs, or a

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Y. Li et al. / Acta Biomaterialia 95 (2019) 73–90 Table 2 Commonly used affinity tags for protein purification in affinity chromatography. Tag

Fusion/Recognition Partner

Function/Purpose

References

His-tag GST FLAG Hemagglutinin antigen

Metal chelate GST-fusion protein M1, M2, M5 mAbs 12 CA5 mAb

[58–60] [61,62] [62–64] [63,65]

Heme tag

Various peptide motifs

S1v1

Various peptide motifs

Streptag II, Twin-Strep-tag MBP

Strep-Tactin protein Amylose (maltose analog)

Self-cleaving intein

Depends on the affinity tag intein is fused with (eg. chitin binding domain)

Purification under native and denaturing conditions Enhances purification and stability with mild elution conditions Calcium-dependent binding reaction for mAb purification Mostly used as a detection tool for recombinant proteins through immunoblotting assays High affinity for specific amino acid sequences found in various proteins (e.g. AzHm14 and MBP-Hm16) utilizing coordination chemistry Multifunctional tag that is a substitute for the His-tag; simultaneously enhances expression, thermostability, and production of recombinant proteins Matrix modified with streptavidin for improved affinity Single step purification via cross-linked amylose affinity resin; mild conditions for elution of target fusion protein; effective solubilizing agent Intein can be fused to many different affinity tags to induce self-cleavage and avoid the risk of nonspecific cleavage of tags and target proteins from proteolytic enzymes

combination of these comprised of synthetic oligonucleotides that can fold into 3D structures capable of binding to a target molecule with high affinity and specificity [44]. Although both aptamer ligands and antibodies can display high specificities for a given protein epitope, aptamers have a distinct advantage over antibodies due to their stabilities in harsh chemical and physical conditions. For example, aptamers can recover their native conformation and effectively bind to target proteins after re-annealing and have been shown to be compatible with stringent caustic cleaning conditions during the separation process [43,55]. Additionally, aptamers can be easily modified to strengthen their insensitivity toward proteases commonly present in biological extracts, as opposed to polypeptide ligands which are prone to enzymatic degradation [46]. Recently, Forier et al. used DNA aptamer affinity ligands to selectively purify human plasma-related proteins from various origins [43]. Three human therapeutic proteins, Factor VII, Factor H, and Factor IX were purified in a single chromatographic step using three distinct aptamer affinity ligands. Factor VII, a protein responsible for blood coagulation in the body, is of very low abundance representing less than 0.001% of total plasma proteins [43]. A DNA aptamer called Mapt2.2CS was used to selectively adsorb Factor VII proteins in the presence of divalent cations and eluted with an aqueous EDTA buffer solution. The binding capacity was estimated to be 4–6 times higher for the aptamer affinity system compared to the standard specific antibody ligand purification system for this protein, with a purity of over 98% [43]. Similarly, antiFactor H aptamer, called MaptH1.1CSO, was used to purify Factor H, a large, low-abundance plasma glycoprotein responsible for regulating the alternative pathway of the complement system, ensuring that immune responses do not damage host cells and tissues [56]. Lastly, Nonapta5.1 aptamer was used in the purification of human coagulation Factor IX, responsible for blood clotting and commonly used in patients suffering from hemophilia B [57]. After Factor IX was selectively captured and washed with 50 mM TrisHCl at pH 7.4 and 2 M sodium chloride, the captured target proteins were eluted with EDTA buffer at pH 8. SDS-PAGE analysis confirmed that almost all of the Factor IX proteins were eluted from the column with perfectly preserved biological activity [43]. 2.2. Fusion tags Affinity tags have also gained widespread attention in the purification of therapeutic proteins. They are comprised of exogenous amino acid sequences with a high affinity for certain biological or chemical ligands. Affinity tags are normally fused to the Nterminus or C-terminus of a target protein, which allows the protein to be selectively captured and purified using a tag-specific resin via affinity chromatography or other methods [72]. Affinity

[66] [67] [68] [69] [70,71]

tags are advantageous to utilize in the downstream processing of protein therapeutics for several reasons. In many instances, the target protein exists as a version with an affinity tag from prior research stages. Furthermore, the high yields of affinity purification make the use of affinity tags economically favorable. Affinity tags can be both beneficial and harmful for the target protein, and are typically removed through the addition of proteolytic enzymes or more recently, through self-cleaving modules that have been developed over the past several years [60,73]. Table 2 shows a multitude of relevant affinity tags commonly used in affinity chromatography for protein purification; several other tags exist that have been reviewed elsewhere, and will not be discussed here [72,74]. His-tags are one of the most prevalent affinity tags used in purification strategies due to their abundance in many different proteins. Additionally, His-tags do not require a specific protein folding conformation to function and have been used to purify soluble membrane proteins stabilized by detergents or lipids, thus demonstrating their robustness [58–60]. Purification of His-tagged proteins is accomplished by using chelated metal ions as affinity ligands that are complexed with an immobilized chelating agent. The target protein is separated through interactions between the imidazole side chain of histidine and the affinity ligands and is purified using immobilized metal affinity chromatography [74]. Recently, Yang et al. used nickel-nitrilotriacetic acid agarose resins to purify His-tagged SmpB recombinant proteins from Mycobacterium tuberculosis [75]. The recombinant proteins with His-tags showed higher soluble protein products compared to glutathione S-transferase (GST), GB1, and thioredoxin fusion tags, with 99% purity and a recovery yield of 26.9 mg/L [75]. Despite their robustness and widespread use, His-tags come with several drawbacks. For instance, there is a propensity for other proteins with external His-residues to contaminate and co-purify with the His-tagged target protein, which has led to the development of the LOBSTR E. coli strain to eliminate the majority of abundant contaminant host proteins [72,76]. Furthermore, in certain cases Histag residues may interfere with target protein folding and activity [77–79]. As a result, various other epitope tags have been developed [72]. One very common epitope tag used for affinity protein purification is the FLAG tag, which consists of a short, hydrophilic octapeptide (DYKDDDDK) [63]. The FLAG tag contains an intrinsic enterokinase cleavage site at the C-terminus, allowing it to be easily removed from the target protein. FLAG is recognized by M1, M2, and M5 mAbs and can be purified using an immobilized mAb matrix under non-denaturing conditions and eluted by lowering the pH or adding chelating agents like ethylenediaminetetraacetic acid [63]. FLAG-based affinity systems have been used to purify Oct4-interacting proteins from mouse embryonic stem

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cells (ESCs) [64]. Additionally, FLAG tags have been used for optimizing signal intensities of enzymes for analysis [62]. For example, Li et al. fused a FLAG tag to Nano-luciferase, an enzyme responsible for bioluminescence, to remove background signals generated from other residual reagents present in bioluminescence assays [62]. Similarly, hemagglutinin tags are commonly used for detection of proteins of interest in cell biology and biochemistry analysis [65]. Another small peptide fusion tag is the heme tag, which binds heme covalently to a protein in E. coli for visual tracking of protein expression and purification [66]. Asher et al. reported an affinitybased purification strategy for proteins expressed in E. coli that utilizes coordination chemistry of a heme tag to an L-histidineimmobilized Sepharose resin [66]. They demonstrated the viability of their platform on two model proteins, Pseudomonas aeruginosa azurin (Az) and E. coli maltose-binding protein (MBP) by purifying them from cell lysate under mild conditions [66]. Recently, in an effort to find superior substitutes for His-tags, a multifunctional tag, S1v1, was created from a self-assembling amphiphilic peptide derived from the Zuotin protein sequence by replacing lysine with histidine residues [67]. The expression levels of three different proteins: polygalacturonate lyase (PGL), lipoxygenase (LOX), and green fluorescent protein (GFP), were monitored after fusion with S1v1. Although protein expression levels were negligible when using His-tags in combination with a PT-linker, the PT-linked S1v1 fusion proteins demonstrated enhanced expressions and bioactivity for all three proteins. The enhanced bioactivity of these proteins can be explained by fusing S1v1 at the Nterminus through the PT-linker, thus leaving the active site at the C-terminus exposed. Such high expression levels are not shown in His-tag fusions because of their fusion to the C-terminus. Additionally, S1v1 fusion proteins were purified using nickel affinity column chromatography yielding high purities and about 47% recovery while His-tag fusion of GFP achieved only 8.23% recovery. PGL and LOX fused with His-tag could not adsorb to the nickel column [67]. New approaches have been made to improve the efficacy of other common affinity tags as well. For example, Schmidt et al. developed Twin-Strep-tag (TST) consisting of two Strep-tag II moieties connected with a short linker [68]. Strep-tag II is advantageous compared to His-tags due to its inertness, resistance to proteases, and independence from metal ions during purification; however, it is not suitable in denaturing conditions and can dissociate in mild buffer conditions in the presence of D-biotin [63]. Such limitations prompted the creation of TST by Schmidt et al., who have utilized the synergistic effect of two Strep-tag II moieties to promote steady, non-competitive high affinity binding with biotin derivatives [68]. This synergistic effect is reversed upon addition of a competitive agent which facilitates elution by blocking the binding site and preventing re-binding. This system is especially advantageous for affinity purification of target proteins from diluted extracts like mammalian cell culture supernatants [68]. In addition to monoclonal antibodies, transcription factors, and enzymes, affinity tags can be used to selectively purify growth factors and optimize their expression and solubility. Recently, human fibroblast growth factor 21 (hFGF21), an essential glucose and lipid metabolism regulator, was fused with MBP in addition to several different affinity tags to compare the efficacy of each tag on hFGF21 expression and stability [69]. Although MBP-fused hFGF21 had solubilities similar to those of other tags such as GST and hexahistidine (His6) at 18 °C, the MBP-tagged protein showed significantly improved solubility in cell culture conditions at 37 °C [69]. MBP-hFGF21 was therefore subsequently purified using affinity chromatography and completely eluted from the column with 1 M imidazole-containing buffer. The MBP tag was removed using Tobacco Etch Virus protease, yielding 8.1 mg of purified hFGF21 as the final product obtained from 500 mL of starting culture [69]. MBP affinity tags have found applications in enhancing cytokine

expression as well. Do et al. employed MBP solubilizing tags to enhance the expression of interleukin-33 (IL-33) in E. coli [69]. IL-33 is an essential alarmin in the immune system which serves to activate many immune cells such as macrophages and natural killer cells upon pathogen exposure or tissue damage [80,81]. MBP-fused human IL-33 demonstrated greater solubility and expression levels in E. coli over a wide temperature range compared to octa(poly)histidine (His8) and GST tagged-fusion proteins and resulted in a final product of 96% purity [69]. A major limitation of using affinity tags for protein purification is the difficulty in tag removal. The predominant method for affinity tag removal is proteolytic cleavage [82], however, this procedure may result in nonspecific cleavage of the tag and/or target protein. Recently, self-cleaving intein tags have been used in combination with other common affinity tags for chromatographybased protein purification to overcome these limitations. Intein tags can be induced to self-cleave through the addition of thiol compounds or changes in pH and temperature. Wood and coworkers demonstrated a system combining intein with a chitin-binding domain (CBD) tag to form a self-cleavable tag capable of purifying various recombinant proteins expressed in E. coli [71]. The high binding affinity of the CBD tag yields a highly purified target protein, which is eluted from the column after intein cleavage upon a change in elution pH [71]. Several other self-cleaving fusion tags have been reviewed elsewhere, and will not be discussed here [60]. Affinity tags present a viable alternative to affinity ligands for enhancing the purification of therapeutic proteins. In addition to purification via affinity chromatography, affinity tags have applications in the isolation and capture of various hormones, cytokines, and other biological moieties for detection and analysis purposes. Over the next several decades, we can expect that the number of affinity tags will continue to rise as newer tags continue to be developed to improve purification strategies for a vast array of therapeutics. 3. Non-affinity chromatography Non-affinity chromatography is commonly used during the polishing steps in the downstream processing of therapeutic proteins. In a typical platform operation for mAb purification, one or two polishing steps follow a protein A affinity capture step to remove residual host cell protein (HCP), DNA, viruses and other productrelated impurities [83]. Unlike affinity chromatography, which relies on the likeness or specificity of an antibody binding to a stationary phase, non-affinity chromatography utilizes charge, hydrophobicity, or molecule size for purification [83,84]. The most frequently used non-affinity chromatographic modes are IEX, either anion exchange or cation exchange, and HIC. Other methods include mixed-mode chromatography (charge and hydrophobicity based separation), size-exclusion chromatography and reversephase chromatography. Current IEX resins utilize a cross-linked matrix commonly made of agarose, silica, polymethacrylate, poly(styrene-divinylbenzene) with a functional ligand, such as a sulfonate group (cation exchanger) or a quaternary amine (anion exchanger), to facilitate binding. HIC resins exploit similar support matrices but have hydrophobic ligands such as ethyl, propyl, or benzyl/aromatic groups to enable adsorption. Non-affinity chromatography is crucial for obtaining high purity in biopharmaceuticals manufacturing. This portion of the review focuses primarily on ion exchange and hydrophobic interaction modalities and mechanisms. 3.1. Ion exchange modalities Ion exchange chromatography has served as an essential tool for the separation of biomolecules for decades and remains a frontrunner in impurity-removal steps and charge heterogeneity

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evaluation [85]. This modality utilizes a charged (negative or positive) stationary phase with immobilized ligands and binds therapeutic proteins under specific solution conditions, particularly pH. With increasing pH, the overall charge of the protein changes from net positive to net negative, with the transition occurring at the pI, the point of zero net charge [86]. Ion exchangers typically operate in one of two modes: (1) bind-and-elute mode where proteins bind to a resin and impurities are forced out, or (2) flowthrough where impurities bind to the resin and the desired protein is allowed to flow through the column. While IEX is one of the most widely-used chromatographic modes for protein purification, it has several limitations as an analytical tool in applications such as biologics characterization. These drawbacks include relatively poor kinetic performance and incompatibility with mass spectrometry instruments due to the use of non-volatile salts [85]. Also, packed ion-exchange beds with porous stationary phases could induce product aggregation and present significant diffusion limitations [87–92]. Another significant drawback of IEX is column or membrane fouling, which decreases efficiency and increases cost over time [93]. These limitations support the development of new technologies to increase the speed, sensitivity, and resolution of separations. Designs include monolithic columns, smaller diameter particles in packed beds, and 2D-liquid chromatography setups [94]. New biomaterials are also being investigated as effective alternatives to traditional ion exchange resins and modalities. 3.1.1. Zwitterionic cellulose beads Zwitterionic polymers have been investigated for several biomedical applications because of their versatile properties and dual-charged nature. They have been used for early cancer detection and can help distinguish between cancerous cells and benign cells [95]. Zwitterionic hydrogels have an ultra-low fouling property, suggesting that they could be used for medical implants or scaffolding [96]. The first zwitterionic ion exchange resin was synthesized in 1951 by Stach, which utilized a styrene-vinylchloride copolymer as the base matrix [97]. A need exists for a greener, bio-based zwitterionic ion exchange resin that will reduce cost and improve productivity. Cellulose is a hydrophilic biopolymer and has been used as an ion exchange scaffold for the purification of monoclonal antibodies [98]. It is an ideal candidate because of its repeating structure and ability to be tuned or modified to meet custom chemical property specifications [99]. Trivedi et al. proposed and synthesized zwitterionic cellulose beads via oxidation and coupling chemistry [100]. Pre-treated cellulose pulp was shaped into spheres by spin drop atomization and regeneration. Positively and negatively charged groups were added using coupling reactions and click chemistry [100]. These beads contain carboxylic anions (–COO ) and quaternary ammonium cations (-N+), resulting in an overall net charge of zero. The bead size ranged from 250 mm to 4 mm [100]. Scanning electron microscopy (SEM) revealed that the zwitterionic beads had macro-pores with thicker walls than non-functionalized beads [100]. The larger pore size, seen in Fig. 2A, B, attributed to increased hydrophilicity. The introduction of ions provides an opportunity to carry proteins and other bio-macromolecules. This suggests that zwitterionic beads could serve as an effective alternative to current, non-renewable ion exchange resins. 3.1.2. Electrospun cellulose nanofibers Electrospinning polymers into layered matrices or nanofibers has recently gained considerable attention because of its capability and feasibility in a wide range of applications [102]. Electrospun nanofiber mats have notable characteristics such as high surface areas, porous structures in the nano-size range, and modifiable structures [103]. This technology was developed and patented by Formhals in the late 1930’s and has seen modifications and

Fig. 2. (A, B) A microscopic view of the Zwitterionic cellulose beads. Reprinted with permission from [100], Copyright Ó 2016, Springer Nature. (C, D) Functionalized electrospun cellulose nanofibers. Reprinted from [101].

improvements throughout the 20th century [104]. Electrospun nanofibers have since been effective in chromatographic column development and protein adsorption [105]. Electrospun cellulose-acetate nanofibers, developed by Liu et al., could replace chromatography methods for polishing steps in downstream processing of therapeutic proteins [106]. Dods et al. worked to improve these nanofibers by adding charged functional groups and compressing various layers into a packed matrix for chromatography applications [101]. A cellulose acetate solution was passed through a charged microneedle and deposited onto a grounded collector as a continuous fiber strand, which formed a non-woven mat-like complex with fiber diameters less than 1 mm. The cellulose acetate nanofiber complex was then cut into small squares, stacked between two aluminum plates, and compressed to form the layered, nanofiber adsorbent structure. Upon compression, heat-treatment, and deacetylation, the cellulose acetate layers were regenerated to a compact cellulose matrix. Diethylaminoethyl (DEAE) ligands or carboxylate (COO) groups were added via alkylation or oxidation, respectively, which act as the charged binding agents [101]. The nanofibers with each of the functional ligands are shown in Fig. 2C, D. Tensile strength of the adsorbents increased with increasing compression pressure and was found to be a critical parameter in the development of a robust nanofiber complex. The increase in mechanical strength suggests that electrospun cellulose nanofibers can be packed into large-scale configurations without significant changes in morphology. Binding capacities were also analyzed, where the COO modified adsorbent had a comparable maximum capacity (47.5 mg/mL) to commercially available packed-bed resins, while the DEAE adsorbent’s capacity (27.4 mg/mL) was significantly less [101]. Bovine serum albumin (BSA) and lysosome were used for DEAE and COO analysis, respectively. Lan et al. developed similar cellulose triacetate (CTA) nanofibers with hydroxyapatite (HAp) functional groups for protein purification [107]. The coupled effect of CTA’s hydrolytic stability and resistance to biodegradation along with HAp’s exceptional

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biocompatibility, high adsorption capacity, and low cost make this system a standout [108,109]. While these nanofibers show potential and have a promising maximum adsorption capacity of 176.04 mg/g, more development and optimization must be completed. With the correct balance of capacity and material, this technology can be tailored to fit a number of bioprocess applications including crucial polishing steps in protein purification. 3.2. Hydrophobic/hydrophilic interaction modalities HIC is exceptional at removing product-related impurities and is an essential tool in downstream mAb purification [110]. HIC works by adsorbing the non-polar sidechains of proteins to hydrophobic ligands on a stationary phase typically in the presence of high salt concentrations and allowing more polar molecules or impurities to pass through. Hydrophilic interaction chromatography (HILIC) works in the opposite manner and utilizes a polar stationary phase with a non-polar mobile phase [111]. Hydrophobic charge-induction chromatography (HCIC) operates similarly but requires no addition of salt to the mobile phase. Desorption is based on electrostatic charge repulsion and is achieved by lowering the pH of the mobile phase [112]. These modalities are effective at removing molecular weight variants (aggregates and fragments) and process related impurities, and are often utilized as polishing steps in the purification of therapeutic proteins [113]. Much like IEX, HIC, HILIC, and HCIC are typically operated in a column format and can suffer from diffusion limitations and fouling. Methods such as grafted resins with newly developed ligands and bio-based monoliths have been developed to improve the efficiency and robustness of these modalities [114,115]. Wang et al. developed a highly selective hydrophilic polymeric monolith with choline phosphate functional groups for analyte separation [115]. Choline phosphate molecules are naturally derived and have been used in the development of biopolymers [116]. Monoliths, in general, have less mass transfer resistance and reduced pressure drops when compared to traditional chromatography columns [117]. Polymeric monoliths are also easy to fabricate, have large surface areas, and exhibit excellent loading capacities [115]. The choline phosphate monolith exhibited an abundance of macropores (>100 nm), permitting high flowrates under low backpressure, which is an improvement from traditional packed beds [115]. Both hydrophilic and electrostatic interactions can be attributed to the separation of polar analytes, which suggests that this monolith could serve as a HILIC stationary phase for the separation of therapeutic proteins. A HCIC resin was developed by Liu et al. that features a poly (glycidyl methacrylate)-grafted polymer and a 5aminobenzimidazole (ABI) functional ligand [114]. Polymergrafted resins have recently been seen to improve protein adsorption capacity and rates in both IEX and HCIC applications [118,119]. The ABI functional group was tested at different ligand densities to determine the optimal protein-binding condition. The G-ABI resin with a ligand density of 330 mmol/g exhibited a protein adsorption capacity of 140 mg/g, which is the highest reported for HCIC resins in literature [114]. Polymer grafting and fine-tuning of the ligand-density can increase available binding area and strengthen interactions. These attributes result in high adsorption capacity and dynamic binding capacity, which makes poly(GMA)-grafted HCIC resins a viable modality for large-scale purification of therapeutic proteins. 3.3. Mixed-mode ligands Mixed-mode chromatography (MMC) is a new purification method that utilizes more than one form of interaction between the stationary and mobile phases. Mixed-mode resins typically

have ligands with multiple functional groups to facilitate hydrophobic interactions, electrostatic interactions, and even hydrogen bonding [120]. MMC resins usually combine ion exchange and hydrophobic interactions to achieve high selectivity and sensitivity [121]. Over the past decade MMC has shown great performance in the separation of mAbs and therapeutic proteins [10,122]. Some MMC resin types utilize a multimodal ligand with a carboxylic group and an aromatic group to facilitate electrostatic and hydrophobic interactions, respectively. Other commercially available MMC materials contain hydrocarbyl amines or sulphur atoms to enable interactions. A significant benefit to MMC is that protein binding can occur without any feedstock dilution or addition of lyotropic salts [123]. While relatively little groundbreaking work has been done in recent years to incorporate biomaterials into MMC resins for protein purification, this technology is promising in its use as a platform chromatography method. 4. Membrane separation Membrane separation is one of the key systems utilized in therapeutic protein purification and downstream processes, such as depth filtration, ultrafiltration, virus filtration, and sterile filtration. Depth filters are commonly used in downstream processes during the cell culture product harvest and post viral-inactivation steps to remove biomass particulates, precipitates, HCP, DNA, and potentially high molecular weight species in some instances [124,125]. Typically, a depth filter is composed of filter aid, binder, and cellulose fiber. The filter’s effective protein absorption capacity was evaluated by Khanal et al. to gain optimal usage [16]. As conventional depth filters contain naturally-derived diatomaceous earth and potentially beta-glucan leachable material that could affect process variability and endotoxin assays, all-synthetic depth filtration media (e.g., Millistak + Ò HC Pro X0SP) has emerged as the alternative to address these challenges and has shown to improve HCP clearance [126]. Sorting molecular components through optimized and engineered barriers could enhance efficiency and selectivity between the feed and permeate streams [127]. To achieve highly concentrated mAbs intended for a subcutaneous injection, ultrafiltration is the main unit operation on the manufacturing scale. For example, Pall OmegaÒ T-series and Millipore PelliconÒ series offer ultrafiltration membranes with different screen channel types, ultra-low protein binding membrane, and different sizes depending on the required scale. However, there are existing challenges associated with this process, such as increased risk of aggregate formation or membrane fouling affecting selectivity and permeability at high protein concentrations [128,129]. Furthermore, physical properties can alter filtration flux and high viscosities can limit operating pressures [130,131]. Various research groups have developed new biomaterials or membrane modifications to enhance the performance of the ultrafiltration unit operation [132]. Membrane surface modifications such as chemical grafting and physical coating post polymer matrix formation provide options for presenting the membrane surface with various functional groups based on specific applications. On the other hand, bulk blending is another approach for membrane modification by which the functional modifiers of interest are added into the membrane casting solution. These membrane modification methods have been discussed and reviewed extensively [133–135]. To enhance the surface hydrophilicity and construct a more porous ultrafiltration membrane structure, Nie et al. used aramid nanofiber (ANF) as a nanofibrous modifier to improve the performance of two representative membranes, namely polysulfone and polyethersulfone (PES) membranes [136]. The ANF nanofibrous structure showed diameters of approximately 30–40 nm and lengths of several micrometers. Various characterization

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Fig. 3. (A) Schematic illustration of similarly sized protein separations through tunable nanoporous block copolymer membrane. (B) Regular order of the membrane’s surface as shown by SEM. (C) Cross-section SEM image showing the thin top layer and the sponge-like bottom layer. (A) (B) (C) were reprinted with permission from [137], Copyright Ó 2013, American Chemical Society. SEM images of (D) 100 nm PS latex beads and (E) SIV particles following filtration on Cladophora cellulose membrane. Reprinted with permission from [138], Copyright Ó 2014, John Wiley and Sons.

results demonstrated increased hydrophilicity in the ANF modified membrane, which enabled higher water fluxes, higher product retention, and enhanced antifouling properties compared to their native membrane structure. Additionally, the ANF modified membrane showed reduced bacterial adhesion when tested with E. coli and Staphylococcus aureus bacteria, which demonstrated its improved antifouling properties. Most importantly, unlike other existing polymer modifiers, ANF can be easily prepared from Kevlar fibers by dissolving them into dimethyl sulfoxide/potassium hydroxide, which is feasible for large-scale production and applications [136]. Separation of similarly sized proteins has been a challenge due to limited fabrication techniques and biomaterials. In the work reported by Qiu et al., an amphiphilic block co-polymer (PS-bP4VP) was introduced to self-assemble into a thin layer of densely packed, highly ordered cylindrical channels with uniform pore sizes of 34 nm and lengths of 100 nm (Fig. 3A–C) [137]. Compared to commercial membranes with similar pore sizes, the water fluxes obtained in this work were one order of magnitude higher, with more than 3200 L m 2 h 1 bar 1 in the middle pH range of

6–8. By customizing the membrane properties with quaternization, the membrane was able to separate similarly sized proteins based on their respective pIs. In this study, BSA and bovine hemoglobin (BhB) were used as the model proteins as they were nearly identical in molecular weights, except BSA has a lower isoelectric point than BhB, pI of 4.7 and 7.0 respectively. Additionally, the authors found that other parameters such as molecular shape, mobility, and the hydrophilic-to-hydrophobic ratio of amino acids could also affect the protein transport behavior through the nanopores [137]. Overall, this work presents a nanoporous block copolymer membrane with tunable properties that targets high selectivity towards a specific protein of interest and could be optimized for efficient bioseparation applications. To achieve high hydraulic permeability and size sieving performance combined with mechanical stability of the separation membranes, many research groups are exploring the composite membranes which consist of nanoporous size-sieving attached to a macroporous support [139–141]. Yao et al. described a micellederived (MD) composite membrane with high selectivity and low flow-resistance [119]. It consists of an asymmetric polystyrene-bl

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ock-poly(2-vinylpyridine) (PS-b-P2VP) nanoporous block copolymer (BCP) size-sieving layer of a few hundreds of nanometers in thickness and a few tens of nanometers in pore size, connected to the PES with a nominal pore diameter of 450 nm as the macroporous support. To bridge the BCP onto the PES membrane, inorganic nanostrands were utilized. Through the swelling-induced pore generation in ethanol, nanopores mean width was estimated to be 20 ± 4 nm and displayed narrow channel-like shape in SEM [119]. Different swelling duration and temperatures were studied to determine the optimal pure water flux and BSA retention values. The authors found the MD composite membrane exhibits 5–10 times higher permeability than commercial UF membrane and demonstrated efficient separation of small globular proteins such as BSA (MW of 67 kDa) and cytochrome C (MW of 14 kDa) [119]. This synthetic assembly system with tailor-made design options allows for different applications of ultrafiltration. BCPs have been widely studied to improve the permeability and selectivity of ultrafiltration (UF) membranes [142–144]. However, tuning the pore size has been a challenge in the field. By controlling the solvent swelling time of PMMA microdomains in the PS-bPMMA films, Ahn et al. demonstrated that UF membrane with nanopores of 6 nm to 17.5 nm in diameter can be generated [145]. The results showed high selectivity and rejection for gold nanoparticles of 20 nm in diameter [145]. With its excellent mechanical stability and promising scalability up to 4-inch diameter nanoporous membranes, this work indicates potential for industrial applications. Furthermore, Shevate et al. described a multi-step preparation to obtain a hierarchical polystyrene nanoporous membrane with a tunable pore size of 6.5 nm to 29.4 nm [146]. The key factor to control the pore size is the solvent exposure time, which dictates the degree of P4VP degradation and transformation in the membrane. The polystyrene membrane with higher hydrophobicity has shown to have efficient separation of emulsion with no oil droplets observed in the filtrate samples. The results also show that the system was able to separate IgG from BSA that has only a 2 times difference in MW [146]. Viral filtration is an expensive operation and generally produced by a synthetic polymer for the biotechnology industry to remove adventitious virus particles from the therapeutic drug product. Metreveli et al. described a natural, unmodified nanofibrous polymer-based membrane, utilizing size-exclusion principles, that was able to remove nano-sized virus/particles by 6.3

log reduction value, which is comparable to industrial filters (Fig. 3D,E) [138]. The nanocellulose filter paper had an average pore size of 19 nm and was shown to efficiently filter beads ranging from 30 nm to 500 nm [138]. With the filter’s ease of largescale manufacturing, it showed potential for applications in the industrial sector. Membranes with nanochannels that feature charged groups can be applied to the purification of biomolecules by allowing the passage of uncharged or oppositely charged solutes, but opposing the passage of co-ions [147]. Sadeghi et al. have developed membranes that feature a packed array of self-assembled micelles with carboxylate functional surfaces [147]. The interstices (1–3 nm minimum) serve as charged nanochannels that allow the passage of solutes. The micelles are formed by self-assembly of an amphiphilic copolymer, poly(trifluoroethyl methacrylate-randommethacrylic acid) [148]. The membranes have been shown to effectively retain negatively charged molecules and allow positively charged and neutral molecules to pass through. In filtration experiments, anionic and neutral compounds were separated with water fluxes comparable to existing commercial membranes [147]. These membranes are also manufactured using easier and more scalable than other comparable approaches. The self-assembled micelle membranes can be used for processes that involve the separation of charged particles, such as IEX in the processing of therapeutic proteins. 5. Magnetic separation Alternative methods for immobilizing affinity ligands on stationary phases have been extensively studied to improve the limited resin capacity of the traditional chromatography methods. Magnetic nanoparticles (MNPs) show great potential in this regard for cost reduction and process integration [149]. The superparamagnetic ability of MNPs can reduce the high cost associated with conventional industrial protein purification such as chromatography, centrifugation, filtration and membrane separation [17,150]. However, to stabilize the particles and to prime the MNPs for therapeutic protein separation, surface modifications such as biomaterial coatings and affinity ligand conjugation are necessary. Combining the possibility of process integration, decreased processing time, and high binding capacity, functionalized MNPs showed substantial potential for downstream protein separation.

Fig. 4. Schematic examples of magnetic nanoparticles. Various coatings and ligands help stabilize and functionalize the magnetic particles for targeted therapeutic protein separation and purification. (A) Dextran coated magnetic particles with a biomimicking ligand for IgG purification. Reprinted with permission from [152], Copyright Ó 2012, American Chemical Society. (B) Molecular imprinted silica coated magnetic particles for hemoglobin separation. Reprinted with permission from [156], Copyright Ó 2013, Royal Society of Chemistry.

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Polysaccharides such as agarose, chitosan, starch, gum Arabic and dextran are suitable coating materials for MNPs. Batalha et al. immobilized affinity ligand 22/8, a protein A mimicking ligand, onto iron oxide MNPs coated with gum Arabic [151]. Similarly, Santana et al. synthesized dextran coated magnetic particles and immobilized them with ligand 22/8 (Fig. 4A) [152]. MNPs’ high surface area to volume ratio increases their tendency to form agglomerates [17,149–151]. The steric repulsion forces created by conjugating these polysaccharides surface coatings help balance the magnetic and van der Waals attractive forces acting on the MNPs, thus stabilizing the particles and mitigating aggregation [149,150,153]. Not only do these biopolymers increase biocompatibility and colloidal stability, but they also are renewable, nontoxic, and biodegradable [150,152]. Additionally, MP_Dex_22/8 was found to have 30 times higher theoretical maximum binding capacity for IgGs than protein A immobilized on agarose used for traditional chromatography [152,154]. Fernandes et al. immobilized Affitins onto their dextran coated MNPs [155]. The antilysozyme and anti-IgG Affitins-immobilized MNPs presented high yield (>95%) of lysosome and IgG separation [155]. While these MNPs demonstrate high binding potential, the iron oxide core can be oxidized under acidic conditions, so binding and elution conditions must be considered [150,152]. With a polysaccharide coating for nanoparticle stabilization and synthetic affinity ligands for therapeutic protein binding, these MNPs show great potential for a one-step recovery of IgGs. Another common MNP surface modification coating is silica. The electrostatic repulsion of the silica shell stabilizes the magnetic core [153]. Further, silica coating also prevents undesired interactions by creating a barrier between the magnetic core and the additional functional agents, such as affinity ligands, linked to the silica surface [149,151]. The advantages of silica coatings lie in their aqueous condition stability, simple surface modification, and tunable surface thickness [151]. Salimi et al. immobilized protein A and protein G affinity ligands onto porous-SiO2 stabilized magnetic particles for IgG separation [157]. They found that the protein A attached-magnetic microspheres have a faster response time and comparable IgG purity and yield to commercial protein A affinity adsorbents used in chromatography [157]. Additional polymer coatings on silica-magnetic particles are also common. Jia et al. synthesized a polydopamine film layer on silica coated MNPs (Fig. 4B) [156]. These particles are molecularly imprinted with bovine hemoglobin as the template protein. Similarly, Li et al. polymerized temperature sensitive monomer Nisopropylacrylamide, functional monomer methacrylic acid, and cross-linking agent N,N’-methylenebisacrylamide on the surface of SiO2 coated magnetic particles [158]. Their surface imprinted MNPs are thermosensitive, allowing them to control adsorption and desorption of BSA simply by adjusting the temperature [158]. While silica coating provides stability to the magnetic core, silica coating with uniform thickness in the nanometer scale still remains challenging [153]. Moreover, imprinted MNPs need improvement on imprinting efficiency and specific template proteins selectivity [156,158]. Surface functionalized magnetic particles show high affinity binding to various therapeutic proteins, such as mAbs, enzymes, lysosomes, and hemoglobin, which can be easily purified. With the versatility of surface modifications, not only can magnetic particles be used for therapeutic protein separations, but they can also be functionalized for cell and nucleic acid separations. Furthermore, various lab-scale autonomous magnetic separators for protein, cell, and nucleic acid separation are already in the market for research and clinical work. However, there is still significant work to be done for large scale purification using MNPs since there are concerns about reusability and product sterility [149]. Nonetheless, magnetic separation’s potential for simplified down-

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stream procedure, high throughput and high efficiency protein purifications, and cost reduction make it a worthy alternative for therapeutic protein separation. 6. Precipitation Protein precipitation has long been developed for protein purification and concentration in both laboratory protein isolation and downstream processing [159]. Ammonium sulfate precipitation is often performed before the chromatography steps to concentrate target proteins and remove impurities based on solubility differences in proteins. However, relying on bulky and expensive centrifuge systems and relatively low selectivity limits its applications in large-scale purification [11]. Nevertheless, methods based on solid-liquid phase separation mechanisms, e.g., crystallization and precipitation, have seen a renaissance in protein purification to achieve high throughput and reduced costs [160]. To overcome the obstacles of current precipitation methods, continuous efforts have been made to develop more advanced precipitants using biomaterials to reduce or avoid the use of column chromatography. Meanwhile, the improvement of membrane filtration techniques will facilitate the wide implementation of the precipitation methods for protein purification. 6.1. Non-affinity precipitation 6.1.1. PEG-induced precipitation Polymers such as polyethylene glycol (PEG) can be used as nonspecific precipitants. PEG precipitation has received broad interest in the field of mAb purification because of its simple operation in mild conditions, fast precipitation kinetics, and low cost. It was originally found that the logarithm of protein solubility follows a linear relationship with the concentration of PEG and higher molecular weight PEGs show higher precipitation efficiency due to excluded volume effects [161]. Linear PEG4000-6000 of high concentrations were traditionally employed and followed by the exploration of branched PEG to reduce viscosity caused by the large linear PEG chains, although protein precipitation yield can be compromised using branched PEG [162]. Recent studies revealed that the efficiency of PEG precipitant could be further characterized by the hydrodynamic radius of PEG, rh,PEG, which takes into account the effects of PEG branching and environmental conditions [162,163]. Conditions such as PEG and protein concentration, pH, ionic strength, and time should be optimized to achieve the best precipitation efficiency. However, PEG is insufficient at removing product-related high molecular weight impurities [164]. Low selectivity remains the major limitation of this technique. In recent years, the combination of PEG with other precipitation methods have been investigated to improve selectivity and achieve higher yields. For example, Oelmeier et al. reported the employment of centrifugal partitioning chromatography (CPC) in combination with PEG precipitation for mAb purification [165]. PEG-driven precipitation was performed in the CPC output stream and results showed that the reduction of HCP was improved from 88.2% to 99.4% after PEG precipitation while recovering 93% of the target protein after re-solubilization [165]. In another study which combined PEG precipitation and chromatography, instead of mAbs, impurities were precipitated by PEG to acceptable levels with the benefit of eliminating the resolubilization step [166]. To address the limitation of PEG precipitation in removing process-/product-related impurities such as DNA and mAb aggregates respectively, the combination of different precipitants, including PEG, caprylic acid, CaCl2, and cold ethanol, were intensively investigated [164,167]. It was demonstrated that the final yield and mAb purity by combining the four

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precipitants could meet drug substance specifications and the caprylic acid/PEG precipitation was competitive with affinity chromatography. In a subsequent study, continuous PEG precipitation of different recombinant antibodies was developed with similar performance to that for batch operation [168]. Recently, the integration of PEG precipitation and cation exchange chromatography was demonstrated a feasible alternative to protein A chromatography [169]. Despite the wide application of PEG-induced precipitation, concerns still remain on the solubility predictions of target proteins in high protein concentrations, the removal of impurity precipitates and PEGs, and the assay interference. Careful investigations and additional validation of results are recommended for the development of PEG-based precipitation conditions [170,171]. 6.1.2. Polyelectrolytes-induced precipitation. Depending on the pH of the environment, therapeutic proteins or impurities can exhibit charged groups and be precipitated by oppositely charged polyelectrolytes due to electrostatic interactions [172]. Extensive efforts have been made to study the mechanism and influencing factors of precipitating proteinpolyelectrolyte complexes [173,174]. Proteins carry positive or negative charges when the solution pH is below or above their pI, respectively. It provides the opportunity for charge-based precipitation of target proteins or impurities by controlling solution pH condition. Typically, anionic polyelectrolytes including polyvinylsulfonic acid, polyacrylic acid, and polystyrenesulfonic acid are used to precipitate mAbs of basic pIs [175]. Conversely, cationic polyelectrolytes such as polyamines can be used to remove negatively charged impurities such as HCP and DNA [176]. The reversible nature of ionization eases the resolubilization of the precipitated complex by tuning the solution pH or ionic strength. Sieberz et al. investigated the impact and interactions of various parameters on the separation of a mAb from a model impurity, BSA, using anionic polyelectrolytes [177]. A Design of Experiments (DOE) strategy was utilized to show that all parameters should be optimized simultaneously to account for any significant parameter interactions [177]. In a later study, influencing factors for the precipitation of BSA with cationic polyelectrolytes were investigated and two statistical models were established using DOE [178]. Recently, poly(glutamic acid) (polyE) was utilized to precipitate IgGs in high-concentration solutions. The formation of reversible liquid droplets of the IgG-polyE complex demonstrated an indispensable role in improving precipitation and redissolution yields to nearly 100% [179]. Future studies in polyelectrolytes-induced precipitations require fundamental

understanding of more complicated systems involving the prediction of 3D structures of proteins, local charge distributions, and multi-type interactions including hydrophobic interactions, hydrogen bonding, etc. [174]. Improvement in the controllable selectivity and reversibility will improve precipitation efficiency and facilitate better recovery performance. 6.1.3. Self-precipitation with fusion tags As mentioned previously, fusion tags (mostly peptides and proteins) have been widely used to purify target proteins through tagspecific affinity chromatography. As increasing attention has been focused on the development of column-free purification methods, a variety of aggregating tags have been explored to induce the selfprecipitation of fusion proteins [19]. Subsequent removal of aggregating tags can be achieved by incorporating a cleavage site or a cleavage tag in favor of chemical cleavage, protease cleavage, or self-cleavage [72]. Mechanisms of how these tags induce the aggregation of proteins can be interpreted by the responsive solubility, high hydrophobicity, and self-assembling behavior of the aggregating tags. In the following section, recent progress in some representative aggregating tags will be discussed. Elastin-like polypeptide (ELP), a representative fusion tag composed of repeating peptides Val-Pro-Gly-Xaa-Gly, where Xaa is any amino acid except for proline, is well-known for its thermally reversible phase transition behavior which has attracted considerable attention in protein purification, drug delivery, and tissue engineering [180]. ELPs in aqueous solutions abruptly aggregate when temperature is raised above the transition temperature (Tt). The pellet can be further redissolved at a temperature below Tt. Meyer and Chilkoti demonstrated for the first time that recombinant proteins can be purified by fusion with thermallyresponsive ELP [180]. No distinguishable change in the specific activity of the thioredoxin–ELP60 fusion protein was observed, even after four thermal transition cycles [180]. A large number of subsequent studies have been reported to optimize properties of ELP-fusion proteins such as molecular weight, Tt, and ELP cleavage to achieve better purification performance for various proteins [181]. A self-cleaving ELP tag was designed by Wood and coworkers to purify RNA polymerase and several recombinant proteins with reasonable yield and purity (Fig. 5A) [182,183]. The tagged target protein could be precipitated with a mild temperature shift and resuspended in a cleaving buffer to remove the ELP tag. A subsequent precipitation step was conducted to separate the cleaved tag from the target protein. A recent study built upon this concept and established a high-throughput platform using 24-well plate

Fig. 5. (A) Schematic of ELP-tagged split intein purification method. Reprinted from [183]. (B) Affinity precipitation of mAbs from cell culture with Z-ELP-E2 nanocages. (1) Mix nanocage stock with clarified culture. (2) Spontaneous aggregation through multivalent crosslinking. (3) Wash pellet by suspending in target wash buffer pH < 5. (4) Elute by suspending pellet in buffer pH < 4. (5) Add salt for selective precipitation of nanocage and collect purified mAb in supernatant. (6) Regenerate nanocage to recycle for future use. Reprinted with permission from [192], Copyright Ó 2017, John Wiley and Sons.

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cultures followed by purification in 96-well plates [184]. It was demonstrated that this high-throughput format showed high well-to-well reproducibility without compromising purity or robustness [184]. Apart from the thermally responsive ELP, many tags were constructed with high hydrophobicity to promote the aggregation of the target protein. These include proteins such as Npro, Ketosteroid isomerase, 4AaCter and short self-assembling peptides like ELK16 and 18A [19]. For example, recombinant cystatin C was expressed as an insoluble fusion protein with a peptide-tag, 4AaCter, for simple and efficient purification steps [185]. It was also found that fusion proteins with 4AaCter-tag could dramatically increase the production level of cystatin C with high immunogenicity and immunoreactivity [185]. Recently, Xu et al. reported recombinant production of influenza hemagglutinin and HIV-1 GP120 antigenic peptides using a cleavable self-aggregating tag comprised of a short self-assembling amphipathic peptide ELK16 (LELELKLKLELELKLK) and an intein molecule. The target proteins were first expressed as insoluble aggregates and released by intein selfcleavage into the soluble fraction with high yield and reasonable purity [186]. In general, using these aggregating tags is a simpler purification method and requires minimal production or processing costs. However, the yield and purity of purified proteins are inferior to that achieved by traditional chromatography methods. Further exploration of more efficient designs of cleavable fusion tags to address co-aggregation of impurities, partial premature cleavage, and incomplete removal of the fusion tags are necessary for more applicable purification systems. 6.2. Affinity precipitation Affinity precipitation colligates the advantages of both specific capture of affinity ligands/tags and reversible precipitation. It seems contradictory to possess both characteristics simultaneously since affinity binding requires thermodynamic stability while precipitation represents a phase separation behavior. In fact, the precipitation of the affinity precipitant and protein complex are often triggered either by environmental stimuli such as temperature, pH, ionic strength, or concentration. This occurs after the formation of the complex or the internal changes in the complex such as crosslinking or aggregation upon the ligand-protein interactions. For the precipitation triggered by external stimuli, the design of affinity precipitants requires one segment to specifically capture target proteins and the other segment to be responsive and reversibly soluble. To render the crosslinking of target protein, a design of one precipitant molecule with multiple ligands is necessary in the case of capturing proteins with more than one binding site. In spite of the strict criteria of affinity precipitants, several types of biomaterials including ELP-based materials, smart polymers, and affinity peptides have been investigated for affinity precipitation. 6.2.1. ELP-mediated precipitation As mentioned above, ELP has been employed as a fusion tag for thermally reversible precipitation. A major drawback of this method is the limited availability of the genes coding for fusion construction especially in the case of newly developed proteins. An alternative method is the creation of an external ELP fused with a binding ligand to capture the target protein and undergo inverse transition cycling. Chen and coworkers pioneered the design of a series of ELP-ligand precipitants for mAb affinity precipitation processing [18,187–189]. The ELP-ligand-protein complex was precipitated at an elevated temperature and/or specific salt concentration at neutral pH. After redissolving the complex, an elution buffer at a lower pH was added to release the target protein. Subsequently, the free ELP-ligands were removed by a second round of thermally

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triggered precipitation. Protein G was first constructed in ELP fusions to purify IgG [187]. Similar to the results with pure antibodies, around 60% and 90% of IgG in the mouse and rabbit serum was recovered, respectively; 100% recovery of the ELP-protein G fusion was achieved in both cases [187]. In addition to the ease of purification and the high efficiency, the avoidance of tag cleavage and favorable reusability property make it a promising method for protein purification [187]. Affinity partners could also be engineered into the target protein and ELP separately in order to customize a binding domain for codable proteins [190]. Recently, ELP fused Z-domain, a shorter synthetic domain derived from the B-domain of protein A, was designed to bypass the lower expression levels previously observed with protein G-fused ELP [188]. High-throughput screening was employed to determine appropriate conditions for the precipitation and elution of mAbs using ELPZ [18,191]. Greater than 99% mAb precipitation yields were obtained at ELP-Z:mAb molar ratio of 4:1 with 0.25 M Na2SO4 for pure mAbs as well as the mAb harvest mixtures [18]. More than 90% overall yields were achieved in elution buffers with pH up to 4.2 [18]. However, the potential denaturation of antibodies at an elevated phase transition temperature and high salt concentration especially in low pH conditions, remain the major concerns for the ELP-Z precipitants. Therefore, exploring new substrates to present antibody-binding ligands at physiological temperature with low/ no salt condition is crucial. Recent progress in creating an ELP-Z functionalized E2 protein nanocage significantly lowered the temperature and salt concentrations due to increased scaffold dimensions and IgG triggered scaffold cross-linking (Fig. 5B) [189,192]. High-yield isothermal purification of IgG without adding salt was achieved in a simple and non-destructive manner. Similar to ELPZ, several other binding ligands such as Z33 and lectin were fused with ELP as the affinity precipitants for purification of human antibodies and glycoproteins [193,194]. With more affinity ligands or affinity pairs being identified, ELP-mediated precipitation has substantial potential for the large-scale downstream processing of therapeutic proteins. 6.2.2. Peptide-based materials and smart polymers. In addition to ELP, several peptide-based materials and smart polymers with affinity ligands have been used in affinity precipitation for protein purification. Peptide-based trivalent haptens were designed with the ability to form cyclic complexes with trastuzumab and rituximab [195]. It was demonstrated that the purified antibodies had native levels of binding to the cells [195]. Unlike ELP-based systems, additional precipitation was not required to remove the trivalent haptens. After the dissociation of the trivalent haptens from the target antibody in specific buffers, membrane filtration can be used to remove the trivalent haptens [195]. This is due to the significant size difference between the trivalent haptens and target proteins [195]. In a recent study, our laboratory proposed to incorporate a protein A mimicking peptide in immunoamphiphiles that can self-assemble into one-dimensional supramolecular nanostructures with a high binding affinity to IgGs [196]. Given the fact that solubility of peptide-based selfassembled nanostructures is highly sensitive to environmental properties such as pH, concentration, and salt conditions, work is being conducted for the development and optimization of this assembled peptide system as an alternative technology for affinity precipitation of therapeutic proteins [197–199]. Smart polymers including thermo-responsive polymers, pH-responsive polymers, and reversibly cross-linked polymer networks are often combined with affinity ligands to create effective precipitants [200–202]. In recent work, a pH-responsive copolymer EudragitÒ S-100 was linked to protein A ligand for precipitation and elution of IgG at pH 5 and pH 2.5, respectively [202]. It was demonstrated that 89% of IgG was recovered from a protein mixture with a purity

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greater than 95%. However, special considerations for pH-sensitive protein-ligand interactions are necessary when designing the pHresponsive polymers. Ideally, the critical pH for protein precipitation and elution should be independent. With the abundant monomer types and easily modulated structures, polymer-based materials show enormous potentials in constructing affinity precipitants. In this section, we discussed the recent advances in developing biomaterials for affinity or non-affinity precipitation of therapeutic proteins in an effort to improve current unit operations and reduce costs in downstream processing. Affinity precipitation is more favorable than non-affinity precipitation due to its high selectivity, however, more efficient methods to remove precipitants or fusion tags need to be developed and validated. Based on existing studies and prior work, several criteria must be satisfied for a successful and efficient precipitant design. First, the precipitant should not compromise the functionality of the target protein and it should be easily removed from the protein solution. Second, high selectivity and high efficiency are needed for either the precipitation of target proteins or impurities. Third, the precipitation behavior should occur in a well-predicted and well-controlled manner. Fourth, the precipitant could be easily manufactured at a large scale and economically. Finally, the precipitant can be conveniently recycled without causing the loss of protein activity. These criteria should be considered for future improvement and development of precipitants for therapeutic protein purification.

7. Aqueous two-phase systems (ATPs) ATPs have been utilized for purification of mAbs, lysozymes, growth hormones, and other therapeutic proteins [203–209]. ATPs utilize the phenomenon of separating two immiscible aqueous solutions under certain critical conditions of concentrations, ionic strength, pH, or temperature [203,204]. Common pairs of aqueous components that can generate biphasic systems include polymerpolymer, polymer-salt, and polymer-ionic liquid [203,205]. The two phases present distinctive densities and are separated into a lower and an upper phase. Therefore, target molecules and impurities can be purified by separating them into two phases of ATPs since different molecules have different partitioning properties for distinct aqueous phases [203–206]. High purity and high yield of therapeutic proteins can be achieved by optimizing the characteristics of the ATPs fluids, such as composition, pH values, ionic strength, and polarity [203–206]. Silva et al. utilized PEG/dextran systems with NaCl to increase ionic strength for mAbs purification directly from serumcontaining cell broth [207]. It was found that IgG’s high molecular weight and hydrophobic surface resulted in IgG preferring the more hydrophobic phase, PEG, while the hydrophilic serum was found in the hydrophilic dextran phase [207]. Furthermore, cells were present at the interface since this would reduce the interface free energy [203,207]. To further improve purity and yield, the researchers then added affinity dual ligands LYTAG-Z to enhance the partition of mAbs towards the choline structural analogue rich-phase (such as PEG and EOPO) [208]. With this system, over 95% clarification and an 89% extraction yield was achieved [208]. To show the viability of large scale protein purification using the ATPs, Schmidt et al. tested IgG purification from cell broth with both continuous and batch systems, and processed up to 100 mL/ min of Chinese Hamster Ovary cell broth [209]. A yield of 80% to nearly 100% from various experimental conditions was obtained, indicating ATPs are compatible for large-scale protein separation and purification [209]. While ATPs have been shown to have efficient clarification and high yield for therapeutic proteins, even in a pilot-scale, there are

still drawbacks. The partition mechanism of the two aqueous phases is not well understood, alongside the complexity for optimizing the two aqueous solutions. Intricate screenings of solution type, concentration, pH, temperature and other parameters all have to be optimized [203,204,206]. Without effective modeling tools for ATPs optimization and the potential high costs of handling and disposal of the phase forming components, ATPs have to be further researched before large scale use for protein purification [204]. 8. Conclusions and future perspectives Therapeutic proteins have attracted considerable attention for the treatment of many diseases. Driven by the rapid expansion of the therapeutic protein market, demands for high-quality proteins will continue to grow in the coming decades at an even faster pace. This puts increasing pressure on biopharmaceutical production especially on downstream processing due to the limited capacities and throughput of established technologies. Enormous efforts have been devoted to improving the productivity and capacity of chromatographic media and to the exploration of single-use or continuous systems. Moreover, debottlenecking the downstream constraints drives innovations and the development of alternative technologies focusing on improving production efficiency and reducing processing costs. These technologies, such as advanced membrane separations, magnetic-based methods, and highselectivity precipitation/phase separations, are currently in the early stages of practical applications to large-scale manufacturing in replacement of traditional chromatography, but may represent a possibility for future adaptations. Most protein therapeutics are delivered intravenously or subcutaneously due to their inherent instability [210], which in some cases limits their medical applications. In addition to developing effective protein delivery strategies, product quality control during manufacturing and drug formulation are equally essential to avoid the presence of product-related as well as process-related impurities. During the manufacturing process, therapeutic proteins may experience a wide range of pH, ionic strength, protein concentrations, and contact materials, leading to the formation of protein aggregates and fragments that might impact the biological activity [211,212]. Multiple strategies have been explored for minimization and removal of these product-related impurities [211,213,214]. In this review, we have described the developments in employing biomaterials in both chromatographic and nonchromatographic techniques for the downstream manufacturing of therapeutic proteins. These materials mainly serve as chromatography media, membrane materials, precipitants, and other absorbents to separate proteins of interest from impurities through affinity binding or utilizing other physical properties such as charge, hydrophobicity, size, differential partitioning, and aggregation propensity. Low production costs, low toxicity, and tunable structures are appealing advantages of biomaterials for protein purification. Recent progress has manifested the significance of utilizing and optimizing these biomaterials for current downstream processing methods as well as for alternative and disruptive technologies. Further development in the design of advanced biomaterials will address the product quality, efficiency, capacity, cost, scalability, and robustness of the new purification systems. We expect these biomaterials to serve as essential elements for disruptive technologies to be developed in downstream processing of therapeutic proteins. Acknowledgement This work was supported by Bristol-Myers Squibb, U.S.A.

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Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi.org/10.1016/j.actbio.2019.03.015.

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