Search from website
Search from website
In this article, we outline the key aspects of cell line development, including host cell selection, genetic engineering and single cell cloning. We also touch on the challenges of productivity, stability, and compliance while highlighting proprietary IcoCell® platform advantages. This overview equips readers with the insights needed to understand the cell line development process, highlighting its key stages and essential details.
A cell line is a defined population of cells that can be maintained in culture for extended periods of time, as opposed to primary cells which have a limited capacity to proliferate. Cell lines are widely used in research, e.g. studying general physiological and pathological processes, production of biological compounds (vaccines, cytokines, hormones, clotting factors, enzymes, therapeutic antibodies) or diagnostics. 29, 56, 57, 68
For industrial manufacturing of biopharmaceuticals, the producer cell line has to be clonal, which means that it is derived from one single cell 3. The so-called monoclonality is required because in inhomogeneous (non-clonal) cell populations, there may be variability in specific productivity and product characteristics between individual cells potentially resulting in quality issues 48, 8. Among other reasons, the variation can be caused by the genomic instability of the cell line, which comes from the plasticity of the genome of the cells, the random integration of the transfected gene of interest into the genome and gene amplification. 43, 51, 76, 84
In most cases the majority of the cells has an average level of protein production, whereas a minority shows a significantly higher productivity 8. High-producing cells use their metabolic resources for protein production and can therefore have lower growth rates, which can cause the cells with lower productivity to outgrow them 48. For that reason, in order to have higher product yields, cell lines have to be derived from a single high-producing cell. It is especially important in the production of therapeutic proteins from a quality and regulatory perspective to eliminate heterogeneity. 
There are many different expression systems available for the production of proteins, including microbial, yeast, plant, insect, mammalian cells. Each expression system has advantages and disadvantages used for specific needs. For instance, microbial expression systems are commonly used for simple proteins because of the low cost, high productivity and fast growth rate of the cell lines, E. coli and S. cerevisiae being the most commonly used 8. For example single chain variable fragments (ScFvs), antibody fragments (Fabs), single-domain antibodies (sdAb), peptide hormones, like insulin and its analogs, and many enzymes are manufactured by using E. coli 21. Although bacterial expression systems are considered as one of the oldest in biotech history, other expression platforms have been developed because of their accumulation of proteins in inclusion bodies, protease contamination from host proteins leading to degradation of the expressed protein, endotoxin accumulation, and, most remarkably, lack of proper post-translational modifications.Yeasts are commonly utilized as host expression systems and are characterized by capability of providing some post-translational modifications, medium to fast growth rate, easy genomic modifications, and low contamination risk 67. Saccharomyces cerevisiae and Pichia pastoris are the most widely used expression systems for biologic manufacturing. Although some yeasts have been genetically modified to carry out human-like N-linked glycosylation, one of the significant drawbacks is the degradation or truncation of the protein of interest, which leads to low yield and loss of functional activity. Insect expression platforms can be used to produce multiple recombinant products, for example the HPV vaccine. However, they lack glycosylation capabilities equivalent to those of higher eukaryotes enzymes 54.However, many biopharmaceutically important proteins, e.g. monoclonal antibodies, require complex folding and post-translational modifications, most commonly glycosylation. That is why mammalian cell cultures continue to be the system of choice for many products such as therapeutic antibodies, recombinant proteins and viral-based vaccines 21.
This article mainly focuses on the process of mammalian cell line development. Some industrially important mammalian cell lines are Chinese hamster ovary (CHO), mouse myeloma line NS0, hybridoma cell lines and human embryonic kidney (HEK) cell line 21. CHO cells are the most commonly used hosts for biopharmaceutical protein production as 70% of biologics and almost all monoclonal antibodies are produced in them 8, 49.CHO cells are preferred because of:they have the ability to grow in suspension culture and therefore can be cultured on a large scale 42, 48are less susceptible to infections by human viruses as few human viruses are able to propagate in them 6, 76can easily be adapted to various serum-free and chemically defined media (which ensures reproducibility between batches) 48, 76 are able to perform post-translational modifications (most importantly glycosylation), which are similar to those in human cells, which make them compatible and functional in humans 42,76.
Icosagen’s proprietary platform IcoCell® serves to efficiently generate stable, biopharmaceutical high-producer cell lines for GMP manufacturing. The IcoCell® is based on fully biosafety tested CHO-S starter cell line, which is stably modified by expression of a specific heterologous expression enhancing nuclear factor, and for an efficient antibiotic-free metabolic selection process. These additional unique elements promote the integration of the transgene into active regions of the CHO genome for high yield and stable protein production.
The cell line development process starts with vector construction and codon optimization. This is followed by the transfection of cells. After transfection, specific selection protocols are applied to pinpoint cells that have been successfully modified. The procedure then progresses to single-cell deposition and the verification of monoclonality, ensuring the uniformity and genetic identity of cell lines. Subsequent stages involve the examination of expression and characterization of bioproducts to confirm their desired properties. The development phase culminates in scaling-up activities and the establishment of master cell banks, which are crucial for transitioning bioproducts from research settings to commercial production. Upstream process development encompasses the series of steps involved in the growth and cultivation of cells within bioreactors, which includes fine-tuning the media and environmental conditions for optimal cell culture. Meanwhile, downstream process development is focused on the purification and processing of bioproducts through methods like filtration and chromatography. 
The process of initiating recombinant protein expression in eukaryotic cells involves the design, construction, and preparation of the DNA expression vector. Subsequently, the DNA sequence, complementary to the gene of interest, is cloned into the expression vector. To increase the expression level of proteins in mammalian cells, codon optimization has been considered as a powerful strategy 74. The genetic code is degenerate, meaning many amino acids are encoded by multiple codons. Organisms favor certain codons, so using less common codons can limit the availability of corresponding tRNAs, slowing recombinant protein production 66. Replacing rare codons with more common ones in the host organism before cloning can improve gene expression 9.
In Icosagen we design an optimized cloning sequence in terms of codon usage, GC content, avoidance of secondary RNA structures and other modifications. We carefully select regulatory elements (promoters, introns, polyadenylation signal) and (a) secretion signal(s) most optimal for particular target protein.
The process of introducing a foreign nucleic acid into an eukaryotic host cell is known as transfection, which can be chemical or physical 46,11. The introduced gene can result in either transient or stable expression. When a gene is transiently transfected, it remains unincorporated into the genome and is only expressed for a limited period of time. Therefore, transiently transfected genetic material is not passed from generation to generation during cell division, and is suitable for quick and high-throughput small scale protein production 71.
For stable cell line development, the genetic material is integrated into the host cell genome, leading to permanent genetic changes that are passed on to future cell progeny 26. This ensures continuous production of the protein of interest over an extended period of time 27.
In Icosagen the linearized DNA expression vector containing all protein coding sequence is transfected by electroporation into the empty IcoCell® CHO host cell line.
The conventional method of transfection has relied on the random integration of expression vectors into the genome 76. That method has two drawbacks: a low efficiency of integration and highly heterogeneous cell clones because of the influence of the site of chromosomal integration (position effect) 4,23,48,84. However, the advantage of this method is a large number of different variants, among which there may be highly favorable ones. Random integration has to be followed by a very time consuming screening of a large number of cells to find the suitable ones from a cell pool. Some methods used for random single-cell transfection are micro-injections and electroporation. 16,34,36 Targeted transgene integration is a more efficient way of performing transfection as it allows the knock-in of gene(s) of interest into well-defined, and transcriptionally active genomic sites. There are several methods for targeted transgene integration, one of the most common ones being the transposon/ transposase system. It improves integration efficiency, increases the stability of the cell line and offers a high titer. 2,73,76 The technology consists of two components: a transposase enzyme and a transposon vector. The transposon carries the genetic information, i.e. the gene of interest and the marker gene. The transposase recognizes the transposon-specific inverted terminal repeat sequences (ITRs) located on both ends of the transposon vector, and transfers the transposon element into the host cell genome via a “cut-and-paste” mechanism 63. There are also other target-specific genome-editing tools, such as Bacterial Artificial Chromosomes (BACs), Ubiquitous Chromatin-opening Elements (UCOEs), zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs) and clustered regularly interspaced short palindromic repeats (CRISPR)- Cas9 system. 41,53,65,76,90 Icosagen’s proprietary DNA expression vector contains unique, built-in DNA cis- elements, responding to the IcoCell® nuclear factor. Firstly, working together with specific sequence motifs on the IcoCell® expression vector, it guides the integration of transgene expression cassette into active regions of the CHO host cells’ chromatin. This ensures reliable and strong expression of the desired recombinant antibody or protein. Secondly, it activates the transgene expression and/or prevents the transgenes from being silenced during the the development process and manufacturing. As a result, the IcoCell® technology ensures sustained, high-level expression of the protein of interest even after many cell divisions in thousands of liters of bioreactor scales.
Illustration of the IcoCell® platform, highlighting the nuclear factor addition and transcriptional activator for enhanced transgene integration and expression.
Transfection can be an error-prone process, because not all cells will successfully receive the transgene. Therefore, using an appropriate selection marker, such as an antibiotic resistance gene, or transgene co-expressed fluorescent protein, is necessary to select and maintain stably transfected cells in culture 13. Additionally, the frequent accumulation of metabolic byproducts like ammonia and lactate in mammalian cell cultures can have a detrimental impact on cell viability and product-specific yield. Therefore, the marker gene can serve a dual purpose: facilitating the selection of stably integrated transgenes and maintaining cell viability and product quality 21. Although antibiotic-based selection systems also exist, this article focuses on two metabolic selection systems.
The glutamine synthetase (GS) system is similar to the methotrexate system, utilizing a specific drug to inhibit an essential enzyme marker crucial for cellular metabolism. The drug used in this system is methionine sulfoximine (MSX), which inhibits GS. The GS enzyme catalyzes the production of glutamine from glutamate and ammonia, which is necessary for the cells to replicate. Cells are transfected with an expression construct containing the GS gene, and grown without glutamine, in the presence of methionine sulfoximine which inhibits endogenous GS activity. An alternative is to use a cell line that lacks the GS gene. Under these conditions, stably transfected cells which express GS can grow, while cells that do not express this protein, die. Currently, cell line development technologies used by most biopharmaceutical companies are based on either the methotrexate (MTX) amplification technology that originated from the 1980s, or glutamine synthetase (GS) system. 48,72 
The IcoCell® host cell line contains further proprietary genetic cellular modifications, allowing for an efficient drug-free metabolic selection process of high-producer clones. Another selection method based on intracellular fluorescence is the methotrexate (MTX) system. The dihydrofolate reductase (DHFR) is a very important enzyme in the cell that catalyzes the conversion of folate to tetrahydrofolate, which is necessary for de-novo synthesis of purines and pyrimidines 28. Fluorescently labeled methotrexate is a drug similar to folate and can bind to DHFR, and inhibit it 88. Without tetrahydrofolate, the cells cannot replicate and die 85. The methotrexate selection system relies on the survival of the cells with sufficient copies of the DHFR gene, as they can synthesize nucleoside precursors, hypoxanthine and thymidine, enabling their survival.
This system can also be used for gene amplification, which is an increase in the recombinant gene copy number in the cells, resulting in higher yield of recombinant protein 48, 72. In order to achieve that, cells are cultured in increasingly higher levels of MTX. Only cells with increased copies of the DHFR gene, and consequently with gene of interest, will survive the selection process 8.
However, the gene amplification process, as mentioned above, leads to heterogeneity in protein expression levels, due to genomic rearrangements 40,43,51. Moreover, multiple rounds of MTX selection require more time for the development of a cell line. Additionally, it’s important to note that increasing the gene’s copy number is only beneficial to a certain extent. While cells selected through this method may contain thousands of genomic copies of the gene of interest, protein levels are typically increased by only 10- to 20-fold. Moreover, having a large number of genomic copies can result in undesired effects, such as reduced stability of the transgenes, position effects, and disruption of other genes. 45, 82, 83
Single cell cloning is performed to ensure that the that the resulting production cell line is monoclonal. The goal of the cloning step is to facilitate the isolation of stable, highly-productive cell populations, and to minimize the genetic and phenotypic diversity within a cell line population 24. This uniformity is essential for the consistent production of biopharmaceuticals, as it ensures that the product will have the same quality and efficacy batch after batch.
The separation of single cells is still a challenging task. Main challenges are the viability, purity of the isolated cells, throughput (in terms of single cells isolated per second), targeted total number of single cells and yield of the method used. Cell viability is required when isolating single-cells for the purpose of production of monoclonal cell cultures, because the cells have to survive the process unchanged in order to be further analyzed, and used to produce the protein of interest. Cells respond to stress factors like mechanical forces or toxic substances, which can lead to differentiation, reduced viability or even apoptosis. The methods have to offer gentle extraction and handling when operating on living cells 31.
Different single-cell cloning methods described in this article include automated cell selection, fluorescence-activated cell sorting, laser capture microdissection, microfluidics, and limiting dilution.
There are also techniques such as single cell printer by Cytena and VIPS (Verified in-situ plate seeding) by Solentim that combine cell seeding with microscopic imaging 76, 10. They offer higher cloning efficiency than traditional methods such as LDC with direct and reliable proof of clonality. Single-cell printing is an improved single-cell cloning method that involves single-cell deposition using Cytena's Single-Cell Printer (SCP) and plate imaging. It detects and analyzes cells in a microfluidic dispenser chip based on cell size and shape. The cells are then ejected within a microdroplet and deposited into microplates. Droplets that have no cell, or more than one cell, are deposited into a waste container. An image of every single-cell ‘printing’ event is created, and can be used as additional evidence that a cell line is monoclonal. 87, 31
Validated In-situ Plate Seeding (VIPS) combines gentle single-cell printing with in-well droplet and whole-cell imaging. It images the cells in the dry empty well and an intelligence-based image analysis then confirms whether a single cell was dispensed or not. If the well contains a single cell, the well is carefully filled with medium. During the clonal growth phase, VIPS uses daily whole-well imaging, recording a timeline of well images and at the same time, performing confluency analysis. Successful clonal outgrowth can so be traced back to a single cell to confirm monoclonality. The advances of this method are the increase in the plate seeding efficiency enabling the use of fewer plates, and the gentle nature of the dispensing process, which results in high viability of the cells. 10, 59 Usually 3 top-performing IcoCell® stable production clone pools are subjected to one round of single-cell cloning. From each of the selected clone pools 96-well plates are seeded with single cells, employing the VIPS™ system (Verified In-Situ Plate Seeding; Advanced Instruments). The VIPS™ system is integrated into Icosagen’s CLD workflow and couples gentle single-cell „printing“ with automated high-resolution imaging. These combined measures provide assurance of clonality from 2 independent steps:
The single-cell status is confirmed by two qualified individuals. Wells which do not contain a single cell are automatically marked and excluded from further analyses.
Automated systems are generally very efficient and have a high throughput 14. These systems, however, can be quite expensive 8.
ClonePix, developed by Genetix, is an automated colony picker that utilizes microfluidics technology and real-time image analysis 48. Cell clones are cultivated in a semi-solid media, which allows the progeny of the single cell to remain as a single colony and limits the diffusion of the secreted recombinant protein. These secreted recombinant proteins are captured by fluorescently labeled antibodies and form a halo structure around the cell colony 8, 48, 51. High producing cells are selected based on the fluorescent intensity of the halo structure. Individual colonies are then picked by micro-pins and transferred to multiwell plates for further characterization 8, 48. The ClonePix system can perform the entire process of imaging 10,000 cell clones and selection of high producer cell clones within an hour, there is less danger of contamination and the system is sensitive enough to isolate rare high producing clones 8,17,48.
The LEAP/ Cell Xpress technology consists of the Cell Xpress software module and the Laser-Enabled Analysis and Processing (LEAP) platform. It uses a process of negative selection to isolate the cells of interest 8. The cells are immobilized and cultured in a matrix. A fluorescently labeled antibody that binds to the recombinant protein is added. Secreted protein surrounding each cell is quantified and the cells of interest are marked. The unwanted cells are killed by a laser 8, 14, 35 . That allows the remaining chosen cells to proliferate and they will finally be transferred to a larger well for further expansion 47, 48.
Besides being fully automated and having a high throughput 48, it is also suitable for fragile cell types and low cloning efficiency cell types and the probability of contamination is reduced as it is a closed system 8, 47 . The main drawback of the system is the possible damage to the high producing cells 8. Automated cell pickers work by using a similar approach, but instead of killing the unchosen cells, an automated pipetting system removes the high-producing cells 14.
The limiting dilution cloning (LDC) is a method where cells are diluted on microtiter plates at an average of less than one cell per well 8, 48. The probability to obtain a certain number of cells per well (0, 1, 2, etc.) is described by Poisson’s distribution 75. Whether wells contain only a single cell has to be confirmed microscopically, because of the statistical nature of the method 8,31. Wells containing single cells are marked and assayed, typically by enzyme-linked immunosorbent assay (ELISA) to determine the protein productivity of the clones. Wells that contain a high product titer are then selected for further growth and analysis. To ensure monoclonality, at least one more round of cloning is recommended, which makes it a quite time-consuming method. 8, 48, 77
This method has traditionally been the most commonly used method to screen for high producer cell clones due to its simplicity (it can be carried out with standard pipetting tools), relatively low cost and gentleness (low effect on cell viability) 31, 70. However, it is time and labor intensive and low-throughput 8, 48. A significant problem is that the probability of achieving a single cell is of statistical nature and there is no way to be entirely certain that the cell line generated is derived from a single cell 77. Therefore clonality is still not guaranteed and further technologies are required downstream to prove the presence of a single cell in a specific well (for example microscopic imaging systems) 31. In order to find a cell line with desired characteristics, tens of thousands of clones need to be evaluated. With limiting dilution, only a few hundred clones can realistically be characterized, thus increasing the chance of missing a high producing clone. This problem can somewhat be relieved by fully automated pipetting robots that can achieve a considerable throughput 8, 31.
Over the years, fluorescence-activated cell sorter (FACS) has been an important method in mammalian cell culture for the production of biopharmaceuticals. The instrument detects cells based on fluorescent markers that enable the isolation of cells based on cell-surface protein expression and uses droplet technology to sort them 15, 39. FACS significantly increased the number of cells that can be screened as it offers a high throughput. Another advantage is the instrument's ability to sort cells based on standardized sorting modes 8, 86. The wide-spread use of FACS systems also makes them accessible to many users 31. The method however has some limitations, such as negative effects on cell viability and high minimal sample volume, preventing the use of rare samples 62. High purity is typically difficult to achieve by FACS systems due to the system consisting of non-disposable components. The limitations of FACS systems strongly depend on the specific instrument, cell type, and application 31.
To sort the cells using FACS, certain biomarkers on cells are tagged by fluorescent markers. Cell suspensions are then lined up by a sheath liquid and pressure driven through a flow cell. The principles of hydrodynamic focusing cause the cells to align. The cell stream passes by a laser beam to provide optical excitation 48. Optical detectors are used downstream to analyze the fluorescent light from the cells. Cell parameters such as granularity and cell size can also be obtained. The collected data is quantified and analyzed 5. The cells can then be sorted by applying a charge to the droplet containing the cell to sort it into a tube or micro well plate. 31, 48 The FACS method cannot be used to detect proteins that are not cell associated so it is not applicable to cell types that secrete its recombinant protein 8. For these types of cells, a mechanism is required to retain the protein of interest in the vicinity of the cell that secretes it 61, 88. Methods such as matrix-based secretion assays and gel microdrop technology can be used in those cases.
To retain the protein of interest in the vicinity of the cell, gel microdrop technology encapsulates the cell in a biotinylated agarose matrix (agarose polymers with biotin attached to them) 1. A specific biotinylated antibody binds to the secreted recombinant protein. Avidin is added and binds the biotin on the antibody and on the agarose, acting as a bridge and linking the protein of interest to the agarose. The protein is detected with a second antibody that is fluorescently labeled. The cell is then further processed using FACS 8, 14, 48. High producing cells are chosen based on fluorescence intensity 8. This method enables the measurement of protein secreted by a single cell and can be used to isolate rare high-producing cells from heterogeneous populations 8, 69. It provides high saturation levels and great restriction on product diffusion 25. However, there are also some drawbacks. Since the single cell occupancy of beads has to be ensured, the cells are seeded with a low density and therefore typically only ∼10–15% of the beads contain single cells. The other limitation is that expertise is needed to effectively use this method 80.
A similar method to the gel microdrop technology is a matrix-based secretion assay. It is based on the cross linking of an artificial matrix on the cell surface in a high-viscosity medium. The cells are then labeled with biotin and either tagged with an antibody that is avidinated or to an antibody that is biotinylated via an avidin bridge. As with gel microdrop technology, the recombinant proteins are detected with fluorescently labeled antibodies and processed with FACS. This method is less time-consuming, but it is not suitable for sorting fragile cell types and requires optimization to a specific cell line. It also remains technically challenging. 8, 38, 60
Laser capture microdissection (LCM) is most commonly used to isolate cells from solid tissue samples 7, 20. With some LCM systems it is possible to dissect living tissue, enabling the extraction of live cells for cell line development 64. The instrument contains optical microscopes and a cutting laser. A tissue section is first observed through a microscope. The target cell is visually identified and the section to be cut off is marked by having a line drawn around it. The laser cuts the tissue along this line and the isolated cell can then be extracted. It is relatively easy to handle and provides a high level of control over single cells, but the throughput is limited by the process being operator based 31. The single-cell integrity might be compromised and purity is not guaranteed (it may remain unclear whether any contaminants were transferred with the single cell) 22,55. There are many different microfluidic devices, but the techniques for isolating single cells mostly fall into main three categories: droplet-in-oil-based isolation, pneumatic membrane valving and hydrodynamic cell traps. Droplet-in-oil-based isolation uses channels filled with oil, and separated aqueous droplets that can be used to contain single cells 31. It has a very high throughput and offers the possibility of ultra-sensitive and rapid assays in comparison to flow cytometry 19, 50. In pneumatic membrane valving, pressurized air is used to close a microfluidic channel by membrane deflection, which traps the cell. The throughput of these systems is typically lower than for droplet-in-oil isolation 31, 30. Hydrodynamic cell traps are structures in a microfluidic channel that allow only one cell to enter the trap as the trap size is adjusted to the average cell size in a given sample. There can be a large number of traps in parallel to one another, which allows a high number of cells to be isolated at the same time 18. Hydrodynamic trapping can be integrated into handheld pipettes 91.
The advantage of microfluidic systems is that they can be operated with very low volumes (cell sample as well as reagents), which makes it possible to use these systems for rare cell samples. Microfluidic systems can also be used as disposables, which minimizes the issue of contamination. A disadvantage is the low degree of flexibility as a specific microfluidic chip is often restricted to one single application. These systems have potential but are still lacking widespread commercial use 31. Icosagen’s proprietary IcoCell® cell line development platform generates stable high-producer clones for large-scale GMP manufacturing of therapeutic protein drug candidates, with competitive titers, timelines and monoclonality data. Icosagen’s CRDMO concept uniquely combines discovery, pre-clinical, and clinical development capabilities, based on strong optimization, analytical and bioprocessing expertise. This uniquely de-risks and speeds up our clients’ biopharmaceutical development projects.
Employing our proprietary, genetically optimized, transcriptionally activated IcoCell®, the gene of interest is integrated into transcriptionally active chromatin regions. This results in stable clone pools with up to 10 g/L titers in non-optimized shake flask production runs. Early material from those pools in grams-scale can be used to start downstream process, analytical and formulation development, stability testing and early POC productions. This is followed by clonal cell line and process development up to clinical GMP manufacturing.
© 2024 Icosagen
