Mammalian Cell Line Selection Strategies for High-Producers

Darrin Kuystermans and Mohamed Al-Rubeai

Abstract With the increase in mammalian cell-expressed recombinant biotherapeutics, the process of accelerating the selection of generated stable mammalian cell lines is becoming a critical step in cell line development pipelines. The selection process is known to be a major bottleneck in obtaining a cell line from development into manufacturing, but with the current sales of biotherapeutics reaching close to US$125 billion with monoclonal antibodies being more than 50 % of the sales, there is no sign of a decrease in demand and the amount of cell line development projects in the pipeline will keep increasing. This means that more efficient cost effective cell line selection strategies are critical to meet demand for affordable biologics. The current advancements in the cell line selection process has helped in this regard, by reducing some of the labor and time required to reduce heterogeneity and determine clonality of a cell line expressing a quality biotherapeutic protein at the highest specific productivity possible. However, challenges remain in dealing with the sheer volume of cells that need to go through the screening process for the determination of a stable highly productive clone. This chapter will provide a summary of the methodology and strategies employed to select the desired cell lines that meet the demands of the biopharmaceutical manufacturing environment from manual selection to automated systems that aid in the mammalian cell line selection process.

Keywords Single cell cloning • Automation • Cell line selection • Bioprocess development • Biopharmaceuticals • CHO • NS0 • Flow cytometry • LEAP • Mammalian cells • Antibody


The biopharmaceutical industry has seen a rapid growth in therapeutic proteins, with the amount of blockbuster biologics on the market increasing by 65 % from 2006 to 2012 alone. Within those 6 years, for example, the total sales of recombinant biologics had doubled from US$63.8 billion to a staggering US$124.6 billion, with therapeutic antibodies being at the top of the class of biologic products manufactured, increasing from 34.5 % of total biologic sales in 2006 to 51.8 % in 2012 (La Merie Publishing 2013), with the predicted sales of monoclonal antibody (mAb) therapies reaching US$70 billion by 2015 (Chon and Zarbis-Papastoitsis 2011). With this increase in demand for clinical and commercial biologic drug substance manufacturing of mAbs and recombinant proteins through mammalian cell culture production platforms, the need for improved strategies to reduce development time whilst maintaining quality attributes and optimizing yields has been made a priority.

Biopharmaceuticals have employed new technologies to ensure that the mammalian cells selected for large scale manufacturing have the ability to produce recombinant glycoproteins with high efficacy and functionality at optimum expression levels. There are several mammalian cell lines that have been developed as expression hosts, with hybridoma cell lines being one of the earliest; however, for the purpose of biologic therapy production, the hosts have mainly been restricted to a small group of well-classified hosts. The Chinese Hamster Ovary (CHO) cell line (Cockett et al. 1990; Milbrandt et al. 1983) has been the primary host utilized for recombinant biologics production such as monoclonal antibodies with popular CHO cell lineages cultured being CHO-K1, DUXB11, and DG44. Apart from the CHO cell line, other cell lines that are commonly used for the production of recombinant proteins are mouse myeloma-derived (NS0) (Barnes et al. 2001; Bebbington et al. 1992), baby hamster kidney (BHK) (Carvalhal et al. 2001; Christie and Butler 1999; Geserick et al. 2000; Kirchhoff et al. 1996), and human embryonic kidney (HEK-293) (Baldi et al. 2005; Schlaeger and Christensen 1999). More recently, the human retina-derived (PerC6) (Jones et al. 2003) cell line, which has the ability to secrete high levels of recombinant product whilst having a low gene copy number (Butler 2005), has also joined this group of wellcharacterized hosts for biologics production. From these cell lines, the only nonanchorage-dependent cell is the NS0 cell line, while the others need to undergo an adaptation process to suspension culture. The media used for these cell lines can be either serum-free or chemically-defined protein free media for the purpose of process homogeneity, reproducibility, and safety. Chemically-defined protein free media formulations have become the preferred media (Lee et al. 1999; Zang et al. 1995; Sinacore et al. 1996) due to the high reproducibility and increased definition, which reduces process variability and makes it easier for a regulatory authority to approve the process. Early recombinant mammalian cell culture processes had titers below 1 g/L for antibody production (Birch and Onakunle 2005). With advances in culture media formulations and process technologies (Kuystermans and Al-Rubeai 2011), productivities of 2–5 g/L in 'generic' fed-batch process are now routinely reported with extended culture such as perfusion processes reporting within 10–25 g/L (Kelley 2009; Chon and ZarbisPapastoitsis 2011).

An important aspect of a biopharmaceutical glycoprotein product is the functional biological activity, which is usually directly affected by the glycosylation pattern, including trafficking and folding within the host production system (Scallon et al. 2006; Walsh and Jefferis 2006; Raamsdonk et al. 2001). Early studies first highlighted the importance of glycosylation in therapeutic proteins and stated that glycosylation profiles of proteins should be approached on a case by case basis during the development of a production process due to the profound influence of host cell type on posttranslational processing (Hooker et al. 1999; James et al. 1995, 1996; Jenkins et al. 1996). One of the main reasons that mammalian cells such as NS0 and CHO are utilized for therapeutic protein production is their unique ability to produce proteins with oligosaccharides attached to their serine/threonine (O-linked glycosylation) or asparagine (N-linked glycosylation) residues (Butler 2005; Seth et al. 2006). Glycosylation also goes beyond affecting the functional biological activity since, once secreted, the stability, aggregation, solubility, and immunogenicity of the protein may be affected, and insufficient glycosylation patterns can lead to rapid clearance in vivo (Kobata 1992; Sinclair and Elliott 2005; Willey 1999; Wyss and Wagner 1996; Sethuraman and Stadheim 2006). The use of mammalian cell lines other than human can cause immunogenic reactions in humans. Mouse cells generate Galα1,3-Galβ1,4-GlcNAc residues which are highly immunogenic in humans, due to the presence of α1,3-galactosyltransferase (Butler 2005; Jenkins et al. 1996). Glycosylation can also be affected by the culture environment (Chee Furng et al. 2005; Patel et al. 1992; Raju et al. 2000). Clone-dependent glycosylation is also seen due to processing inconsistencies where glycans can have frequent structural heterogeneities (Patel et al. 1992; Seth et al. 2006; Varki 1998). In addition, the enzymes present in a host cell line, which are required for glycosylation to take place, can be species-dependent. Terminal N-acetyl-neuraminic (NANA) sialylation is predominant in humans but varies in other species which tend to predominantly have N-glycolyl-neuraminic acid (NGNA) rather than NANA (Raju et al. 2000). CHO cells have an advantage in sialylation due to their high percentage of NANA sialylation rather than NGNA sialylation taking place on glycoproteins secreted from the cell line (Baker et al. 2001). To optimize all of these glycosylation patterns, glycosylation engineering may be used as a tool to modify the host production line in order to achieve near-human glycosylation. This may require only slight modifications in cells from mammalian origin other than human, but if the host is not mammalian in origin, it can be a complex task. Hamilton et al. (2006) achieved mammalian glycosylation in Pichia pastoris in order to produce functional recombinant erythropoietin. The yeast Pichia pastoris was able to secrete human glycoproteins with fully complex terminally sialylated N-glycans after removing all yeast glycosylation genes and introducing 14 heterologous human glycosylation genes; however, this technology still requires years of further fine-tuning, development and characterization before being approved as a platform for therapeutic manufacturing. There have been similar attempts for insect-, and plant origin-based cell lines where the re-engineering involved can be complex (Hollister and Jarvis 2001; Cox et al. 2006), but the hurdles that are also required to get approval for such a cell line as a manufacturing platform can take many years.

Recently, the emergence of the ability for targeted genome engineering provided via programmable site-specific nucleases such as zinc finger nucleases (ZFNs) (Kim et al. 1996), transcription activator-like effector nucleases (TALENs) (Boch et al. 2009), and the clustered regularly interspaced short palindromic repeats (CRISPR)-associated endonuclease Cas9 (CRISPR/Cas9) system (Mali et al. 2013a, b), an RNA-guided DNA endonuclease, has opened up further possibilities of cell line development for improving product yields. Given the ability to manipulate any gene or insert sequences of DNA at specific sites within the genome of the cell (Gaj et al. 2013), these programmable nucleases can be a powerful tool in the genetic engineering arsenal, as improvements in avoiding off-target effects are quickly being addressed to improve specificity and efficiency (Carlson et al. 2012; Ran et al. 2013; Mali et al. 2013a; Fu et al. 2014). The use of these genome editing systems is still in its infancy in regards to obtaining high-producer cell lines via genome editing; however, some work has already been done to demonstrate the applicability by engineering CHO cells via zinc finger nuclease-aided deletion of Bax and Bak genes to gain apoptotic-resistant cells (Cost et al. 2010).

The intent to culture mammalian cells as an industrial platform has always had its challenges, but the one seen as far back as the 1950s was the natural heterogeneity of mammalian cells, which was first observed when attempts were made to isolate them (Sato et al. 1957). What may seem to be the same mammalian cells in culture can still be cells with very different physiological characteristics and properties, including maximum viable cell numbers, integral viable cell times, specific growth rates and specific production rates of recombinant proteins (Barnes et al. 2001; Kim et al. 1998; Marder et al. 1990; Kim et al. 2001). This inherent heterogeneity seen in culture meant that biopharmaceutical companies have implemented strict procedures to determine 'clonality' in a cell line, in essence meaning that a cell line should be derived from a single cell (ICH Guideline 1995). Cloning procedures are carried out to reduce heterogeneity where a single cell, producing the recombinant protein of interest, is selected from a population of cells in order to ultimately obtain a homogeneous population of cells from a single clone; this population can be utilized for a specific finite amount of generations in the manufacturing environment without any change in expression level in terms of specific productivity, viable biomass and the recombinant product itself.

The selection of high-producing cell lines is known to be a costly process that can be a major bottleneck in getting a cell line from the developmental pipeline into the manufacturing environment. The methods employed to select these highproducers can vary, but the goal is to create a cell line that can be deemed stable due to its consistent product expression and suitable growth characteristics. The stability of a cell line is determined by several regulatory events within the cell, which can be separated into three phases (Barnes and Dickson 2006), where phase one is the vector design, transgene-containing plasmid integration and the chromosomal environment. The second phase is mRNA stability and processing, which is crucial for regulating the expression of transgenes, as is seen with splicing events with XBP-1(S) that has been linked to the control cell line productivity (Shaffer et al. 2004; Sriburi et al. 2004; Yoshida et al. 2006), while the last phase is translational and secretory events where specific proteins within the secretory pathway can affect specific productivity (Smales et al. 2004; Alete et al. 2005). Although several regulatory events within the cell can determine stability, it is primarily assessed by the expressed protein being biochemically comparable, using available analytical techniques, from the initial cell selected to the final cell line produced, whilst observing consistent growth characteristics. Stability in this case can be classified as the limit of the in vitro cell age for production, which itself is a measure of time between vial thaws of selected cells to the harvest process from a bioreactor by measuring the elapsed chronological time in culture via the population doubling level of the cells. Thus, studies to determine cell line stability can be performed by evaluating the secreted quality and expression level of the therapeutic product for a cell line consistently attaining a high maximum viable cell number of over 50–60 generations from the establishment of the Master Cell Bank (MCB) to the end of the commercial manufacturing process in a bioreactor culture.

In order to obtain high-producing cell lines, certain host cell lines have adapted common expression systems that aid in the selection process. Some of these expression systems have become standard in industry, such as the DHFR (dihydrofolate reductase) and GS (Glutamine synthetase) systems. The DHFR system is used with CHO cells (Lucas et al. 1996), where a mutant CHO dhfrcell line is transfected with a vector containing the target gene along with the dhfr marker gene. Selection occurs in hypoxanthine-, glycine-, and thymidine-free media. At the same time, methotrexate (MTX) concentrations are gradually increased to amplify the expression of the protein product due to MTX being able to inhibit the DHFR enzyme (Fig. 11.1). The GS system has been utilized in NS0 (Bebbington et al. 1992) and CHO (Cockett et al. 1990) cell lines, where cells are transfected with a vector containing the gene of interest in addition to the GS gene. Cells possessing the GS gene can synthesize their own intracellular glutamine; thus, these cells can be cultured in glutamine-free media with the addition of glutamate and the ammonia group provided by asparagine (Barnes et al. 2000). Unlike CHO cells, NS0 cells have no endogenous GS activity, making them the preferred cell line for use in this expression system. Selection can be carried out in the presence of methionine sulfoximine (MSX) which inhibits GS activity; as a result, the endogenous GS activity of CHO cells can be circumvented with the use of higher MSX concentrations and by inhibiting the GS activity, the gene of interest is amplified with a gradual increase in MSX concentration. In comparison to the DHFR system, one advantage of the GS selection system is that the cell cultures produce less ammonia, which can negatively affect the glycosylation of the recombinant protein (Yang and Butler 2000a, b); this is in addition to the GS system usually providing a shorter selection period due to the system requiring lower gene copy numbers for a similar expression level of the amplified transgene. Even though both MSX and

Fig. 11.1 MTX cell selection procedure. Illustrates the incremental step increases in MTX concentration and the selection of cell populations resistant to MTX in order to select the few cells with increased dhfr expression. After each round of MTX selection, the cells are exposed to even higher MTX concentrations

MTX selection aid in selection and gene amplification, both methods produce cell clones that are highly heterogeneous with only a few stable high-producer clones. The major cost in the development of biopharmaceutical processes is the time required for cell line development, which traditionally involves the process of gene integration and amplification, with the use of selection agents such as MTX or MSX, followed by single cell selection using limited dilution single cell cloning (LDSCC) techniques. This selection process is laborious and time-consuming in nature (Borth et al. 2000; Carroll and Al-Rubeai 2004; Browne and Al-Rubeai 2007), using manual microscopy techniques to monitor the process and requiring several expansions of possibly hundreds to thousands of clones. In addition, the high-producing clones tend to be overgrown by the low or non-producers due to energy being diverted towards recombinant protein production instead of growth (Al-Rubeai 1999; Borth et al. 2000; Kim and Lee 1999; Kromenaker and Srienc 1994); these tend to be in the majority, since there can be a range of 40–90 % of non-producers, depending on transfection efficiency (Lee et al. 2006). Even after high-producer selection, another round of screening may be required to ensure stability of integration of the transgene before a limited scale-up of a number of selected clones can be carried out to evaluate batch and fed-batch growth profiles. This means that the longest steps in cell line development are the selection procedures, which can take anywhere from 6 to 18 months depending on cell line, expression system and selection method.

With the increase in blockbuster therapeutic glycoprotein approvals for commercial manufacturing, the industry is constantly looking to reduce the cost of marketing these biologics. Cell line development is one area that can greatly benefit from the faster selection of stable high-producers. Although expression technologies, improved media formulations and bioreactor culture systems have resulted in higher yields from smaller more intensified production runs, there is still much development needed in the high-producer cell line selection process in order to reduce labor, development timelines and capital expenditure. This chapter will focus on those methods that are used in cell line selection from single cell screening approaches to recent advances in high-throughput cell screening technologies.

< Prev   CONTENTS   Next >