CELL FACTORY ENGINEERING: CHALLENGES AND OPPORTUNITIES FOR SYNTHETIC BIOLOGY APPLICATIONS

Abstract

The production of high-quality recombinant proteins is critical to maintaining a continuous supply of biopharmaceuticals, such as therapeutic antibodies. Engineering mammalian cell factories presents a number of limitations typically associated with the proteotoxic stress induced upon aberrant accumulation of off-pathway protein folding intermediates, which eventually culminate in the induction of apoptosis. In this review, we will discuss advances in cell engineering and their applications at different hierarchical levels of control of the expression of recombinant proteins, from transcription and translational to post-translational modifications and subcellular trafficking. We also highlight challenges and unique opportunities to apply modern synthetic biology tools to the design of programmable cell factories for improved biomanufacturing of therapeutic proteins.

1. INTRODUCTION

The 1986 FDA approval of recombinant tissue plasminogen activator for acute myocardial infarction and the monoclonal antibody Muromonab for the prevention of kidney transplant rejection marked a milestone for the biopharmaceutical industry and heralded the era of protein therapy. Recombinant proteins currently comprise the fastest-growing sector of the biopharmaceutical industry and are used for the treatment of many conditions including cancer, inflammatory, respiratory, and cardiovascular diseases, and organ transplants. The global market of biopharmaceuticals was estimated at $407.72 billion in 2023 with a compound annual growth rate (CAGR) of approximately 8.03% (Mordor Intelligence, Biopharmaceuticals market analysis - industry report - trends, Size & Share). Twenty out of the 53 new drugs approved in 2019 by the FDA are biologics and include the breakthrough vaccine for Ebola virus infection, and novel drugs for the treatment of cancer and rare diseases (Morrison, 2020). The FDA has mandated provisions for the accelerated approval of drugs for the treatment of cancer and other life-threatening diseases. Furthermore, the introduction of biosimilars is expected to expand the therapeutic protein market to geographic areas currently unable to access blockbuster therapeutic proteins. The increasing rate of approval and introduction to the clinics of protein therapeutics and the consequent rise in production demand has created a pressing need to improve process design and optimization to generate large and economical supplies that meet the market demand.

Monoclonal antibodies are the largest class of biopharmaceuticals and include antibody derivatives such as bispecific antibodies, antibody fragments, and antibody conjugates. Monoclonal antibodies are currently approved for the treatment of a variety of diseases, ranging from rare and orphan indications affecting populations of a few thousand individuals, such as paroxysmal nocturnal hemoglobinuria or the cryopyrin-associated periodic syndromes, to those affecting hundreds of thousands, including some forms of cancer and multiple sclerosis, and millions, such as asthma and rheumatoid arthritis (Hongrong Cai, 2018). Monoclonal antibodies are also extensively employed for the development of in vivo diagnostics and imaging purposes (Leader et al., 2008). The clinical use of recombinant proteins, however, extends beyond antibodies and includes enzymes, vaccines, hormones, and growth factors.

Host systems used for the production of recombinant proteins include bacteria, yeast, insect cells, mammalian cells, and transgenic plants and animals (Tripathi and Shrivastava, 2019). The choice of the host system is typically dictated by the protein-specific post-translational processing and modifications (e.g., glycosylation, disulfide bond formation, phosphorylation, and proteolytic processing) and the cost of production. E. coli cells are widely used for the production of recombinant proteins due to their rapid growth, low costs, and, in most cases, high production yields. Despite the universality of the genetic code, the expression of complex eukaryotic proteins in E. coli cells cannot be taken for granted as bacterial cells cannot support the production of large proteins with complex post-translational modifications, which often accumulate in the form of inclusion bodies (Gupta and Shukla, 2016). In addition, the removal of endotoxin from recombinant therapeutic proteins remains challenging and expensive (Mamat et al., 2015). Yeast cells provide an attractive alternative as they combine the advantages of microbial systems with the ability to reproduce some of the folding features and post-translational modifications characteristic of mammalian proteins. The formation of N-linked and O-linked fungal glycans and hypermannosylated structures, however, often affects the therapeutic properties of the protein product and may elicit an immunogenic response (Tripathi and Shrivastava, 2019; Vieira Gomes et al., 2018). Insect cells have also been used for protein manufacturing due to their high production yield, amenability to scale-up, and ability to carry out more complex folding post-translational modifications compared to yeast cells. The lack of sialylation machinery and tendency to produce immunogenic glycan structures, however, have limited the use of insect cells for the manufacturing of therapeutic proteins (Tripathi and Shrivastava, 2019; Xiao et al., 2014). Recombinant proteins with complex folding and essential post-translational modifications used in biomedical applications are optimally produced using mammalian cells, which present the native protein processing apparatus. Not surprisingly, mammalian cells dominate other host expression systems in terms of the number of FDA-approved therapeutics. Commonly used mammalian host cell lines for recombinant protein expression are Chinese hamster ovary (CHO), human embryonic kidney (HEK 293), mouse myeloma (NS0 and Sp2/0), human fetal retinoblastoma (PER.C6), and baby hamster kidney (BHK) cells (Table 1).

Table 1.

Examples of recently approved biopharmaceuticals with their expression host systems and manufacturers.

Host cell	Product	Manufacturer	Year
CHO cells	Benepali (etanercept)	Samsung Bioepis	2016
	Taltz (ixekizumab)	Eli Lily	2016
	Rebinyn (rh coagulation factor IX)	Novo Nordisk	2017
	Refixia (non-acog beta pegol)	Novo Nordisk	2017
	Lifmior (etarnecept)	Pfizer	2017
	Truxima (Rituximab)	Celltrion	2017
	Tremfya (guselkumab)	Janssen	2017
	Adynovi (rucioctocog alfa pegol)	Baxalta	2018
	Andexxa (coagulation factor Xa)	Portola	2018
	Retacrit (epoetin alfa-epbx)	Eprex and Erypo	2018
	Shingrix (zoster vaccine)	GlaxoSmithKline	2018
	Aimovig (erenumab-aooe)	Amgen	2018
	Zessly (infliximab)	Sandoz	2018
	Herzuma (trastuzumab)	Celltrion	2018
	Enhertu (famtranstuzumab deruxtecan-nxki)	Daiichi-Sankyo /AstraZeneca	2019
	Adakveo (crizanlimab-tmca)	Novartis	2019
	Reblozyl (luspatercept-aamt)	Celgene corporation	2019
	Polivy (polatuzumab vedotin-piiq)	Genentech Inc.	2019
	Skyrizi (risankizumab-rzaa)	AbbVie Inc.	2019
	Evenity (romosozumab-aqqg)	Amgen	2019
HEK	Alprolix (eftrenonacog alfa)	Biogen	2016
	Vihuma (simoctocog alfa)	Octapharma	2017
NS0	Lartruvo (olaratumab)	Eli Lilly	2016
	Trogarzo (ibalizumab-uiyk)	TaiMed/Theratechnologies	2018
BHK	Kovaltry (octocog alfa)	Bayer	2016
PER.C6	Rekeovelle (follitropin delta)	Ferring	2016
Sp2/0	Inflectra (infliximab-dyyb)	Hospira	2016

However, due to the diverse range of posttranslational processing, a universal host that guarantees high yield and quality for all recombinant proteins does not exist. Some proteins are considered “difficult to express” (DTE) even in commonly used mammalian cell lines, such as CHO (Wang et al., 2019). Other eukaryotic expression hosts that can produce correctly folded and fully functional products such as insect cells have been explored as they may provide alternative host systems for DTE proteins.

Protein expression in mammalian cells engages a series of cellular pathways for protein synthesis, folding, post-translational modifications, and subcellular trafficking. Cell engineering strategies aimed at maximizing the expression of recombinant proteins typically overload the native pathways mediating protein folding and processing, often leading to the accumulation of off-pathway protein intermediates and proteotoxic stress (Demain and Vaishnav, 2009; Kafri et al., 2016). The aberrant accumulation of newly synthesized polypeptides and off-pathway misfolding intermediates causes the activation of a series of signaling cascades aimed at restoring homeostasis through the inhibition of global protein synthesis and the upregulation of protein folding pathways in an attempt to ameliorate the cellular protein processing capacity. If sustained, stress leads to the activation of cell death mechanisms, eventually affecting protein productivity due to a decrease in cell viability. These observations point to the critical need to engineer cell factories for enhanced and sustained production of recombinant proteins.

The successful construction of the first synthetic gene circuits in the early 2000s (Elowitz and Leibler, 2000; Gardner et al., 2000) fueled widespread efforts to program biological systems for dynamic, tunable responses, leading to rapid progress in the field of synthetic biology and the development of a toolbox for building computing-like behaviors. Most of the progress has benefited host systems characterized by robustness and ease of manipulation, namely bacteria and lower eukaryotes. Previously characterized genetic parts are used to build programming modules that can be further interconnected to create complex genetic networks controlling a diverse range of functions (Bashor and Collins, 2018). Because these genetic circuits typically display relatively high robustness, the resulting cellular behavior can be recapitulated by mathematical models, which, in turn, provide valuable predictive tools to create new synthetic networks (Ellis et al., 2009). These advances led to defining an engineering-based methodology that has recently generated a toolkit and design rules to build synthetic gene networks in more complex systems such as mammalian cells. As a result, we can now program cells to report on complex pathways with high sensitivity and dynamic resolution (Origel Marmolejo et al., 2020) and control complex processes such as the formation of organoids for drug delivery and regenerative medicine applications (Li et al., 2015). These advances in the field of mammalian synthetic biology offer tremendous opportunities to characterize and control the molecular and cellular mechanisms underlying the production of recombinant proteins and improve the cell factories for biomanufacturing.

Over the past several decades, efforts to meet the increasing demand for high-quality, affordable therapeutics have focused on improving bioprocessing techniques, media optimization, and genetic engineering techniques. Recent advances in synthetic biology provide new tools to reprogram cells for user-defined, precisely controlled biomolecular outputs. In this review, we describe the status of cell engineering for biomanufacturing and discuss the areas of opportunities to apply synthetic biology tools for the design of next-generation cell factories that overcome current limitations to the large-scale production of high-quality protein therapeutics. As most therapeutic proteins are processed through the secretory pathway and secreted extracellularly, we provide a detailed analysis of each step affecting biomanufacturing from cell transgene insertion to protein expression and processing and subcellular trafficking (Figure 1).

Figure 1. Schematic representation of a cell factory of monoclonal antibodies.

Schematic representation of a cell factory of monoclonal antibodies containing a bicistronic transcription unit encoding the heavy and light chains of the monoclonal antibody and a monocistronic transcription unit encoding a chaperone that assists protein folding and processing through the secretory pathway. Overexpression of chaperones prevents the accumulation of misfolded proteins and facilitates three-dimensional folding, resulting in the secretion of natively folded, active products.

2. THE SYNTHETIC BIOLOGY TOOLBOX FOR OPTIMIZING THE EXPRESSION OF RECOMBINANT PROTEINS

The workflow of recombinant protein biomanufacturing involves careful integration of a series of steps affecting the efficiency, quality, purity, and quantity of the protein product. Synthetic biology provides a set of tools uniquely suited to engineer cells for optimizing protein production with tremendous potential for improvement at every step of the workflow, ranging from the delivery of the transgene to the control of transcriptional and translational processes, and optimization of post-translational modifications and secretion processes (Figure 2).

Figure 2. The synthetic biology toolbox for optimizing the expression of recombinant proteins:

(1) Transgene integration tools; (2) Promoters, enhancers, and insulators; (3) Other regulatory elements that enhance protein expression; (4) Regulatory elements that control the expression of proteins with quaternary structure; (5) Engineering translation and trafficking; and (6) Recapitulating native posttranslational modifications. I: Insulator, ENH: Enhancer, LC: Light chain, HC: Heavy chain, IRES: Internal ribosome entry site, WPRE: Woodchuck hepatitis virus post-transcriptional regulatory element, pA: Poly-A tail.

2.1. Transgene integration tools

Therapeutic proteins are produced recombinantly by engineering cells for transient transgene expression or via transgene integration into the host genome. The choice of transgene delivery depends on the features of the protein product as well as the requirements of the production process. Transient delivery via cell transfection is based on an extrachromosomal gene vector which is diluted with each cycle of cell division, resulting in a decrease in protein production over time. Transient transfection reactions require large quantities of DNA and present great batch-to-batch variability in the efficiency of vector uptake. This approach is still typically preferred during the initial stages of process development, as it provides a rapid and inexpensive method to optimize the vector design and culturing conditions. Specifically, transient transfections are amenable to the isolation of highly productive clones in high throughput screens, measurements of in vivo efficacy, and early product development (Aricescu et al., 2006; Gutiérrez-Granados et al., 2018).

Transgene integration into the host cell genome results in more stable and consistent titers of recombinant proteins compared to transient transfection-mediated expression, typically leading to higher yields. The generation of stable cell lines requires the selection of polyclonal populations of transfected cells, which may be followed by a screening of single clones to identify monoclonal populations with the desired characteristics. The method of integration usually determines the number of sites of integration and the specificity of integration, with long-term, stable expression requiring integration at transcriptionally active sites that are not prone to silencing (Mutskov and Felsenfeld, 2004; Richards and Elgin, 2002).

Viral vectors derived from lentivirus and adeno-associated virus (AAV) target active transcription sites in a wide variety of cell types resulting in stable integration of the transgene for a long period of time (Oberbek et al., 2011). Since multiple viral particles can infect each cell, transduction procedures based on a high multiplicity of infection can lead to multiple integrations and high transgene expression levels. The extent of control over transduction and integration efficiency, however, is limited. As a result, viral delivery typically generates highly heterogeneous populations in terms of the yield of recombinant proteins. In addition, because the packaging capacity of lentiviral capsids (~10 kb) limits the insert size, viral transgene delivery is not amenable to the generation of cell lines for the expression of large proteins (Kumar et al., 2001). The safety concerns associated with the use of virus-based systems for the production of protein products intended for human use have further limited this mode of transgene delivery.

Methods of integration based on DNA transposons offer an alternative to viral delivery systems that present an increased level of transgene integration and specificity for transcriptionally active regions of the host genome. DNA transposons are repetitive sequences consisting of a gene flanked by two inverted terminal repeats (ITRs). Transposon systems consist of a transposon donor vector expressing the transgene of interest (up to 60 kb) flanked by the ITRs and a helper plasmid expressing the transposase which mediates the transposition of the transgene from the donor plasmid into the chromosome (Ahmadi et al., 2017; Claeys Bouuaert and Chalmers, 2010). Modulating the ratio between the helper and donor vectors allows optimizing the efficiency of transgene insertion, which, in turn, affects the yield of transgene expression (Ahmadi et al., 2017). The PiggyBac system is a widely used transposon-based integration system. CHO cell lines generated using the PiggyBac system present up to a 4-fold improvement in the volumetric productivity of an IgG1 antibody and stable production yields over a period of 3 months compared to standard transfection methods (Balasubramanian et al., 2015). Transposase technologies have enabled the generation of stable cell factories for the production of therapeutic proteins for clinical use as they increase the homogeneity in comparison to cell lines generated via random transgene integration (Schmieder et al., 2022).

Site-specific recombination methods allow generating highly productive clones through the integration of the transgene at specific integration sites, thus providing control over the chromosomal context and the number of copies of the integrated transgene. Several recombination systems have been used to control the recombination of genetic material, including the bacteriophage P1-derived Cre recombinase, the yeast-derived FLP recombinase, and the lambda phage-derived BxB1 integrase (Bode et al., 2003; Wilson and Kola, 2001). These systems require establishing a cell line with a ‘landing pad’ containing the recombinase target sites. Transgene integration occurs upon co-transfection of the engineered cell line with vectors for the expression of the transgene as well as the recombinase gene. Landing pads containing multiple well-characterized integration sites allow integrating multiple transgene copies at specific sites, enabling stable, long-term transgene expression, and ultimately enhancing the transgene expression levels (Gaidukov et al., 2018; Zhang et al., 2015). Cell lines containing landing pads for transgene integration also enable the development of clones with multiple integration sites for the production of different recombinant proteins (Grav et al., 2018). Integrating multiple transgenes at distinct loci (Baser et al., 2016) that result in different expression levels, on the other hand, could provide a powerful strategy for the co-expression of multiple proteins or protein subunits at the desired relative ratios, which is often a challenging requirement for the production of complex proteins with quaternary structures such as antibodies.

Synthetic biology tools may provide new opportunities to overcome the limitations of traditional integration methods. Targeted transgene insertion, for instance, can be achieved using programmable nucleases such as transcription activator-like effector nucleases (TALENs), zinc finger nucleases (ZFNs), and clustered regularly interspaced short palindromic repeats (CRISPR)-associated (Cas) RNA guided nucleases (Kim and Kim, 2014). These nucleases induce a double-stranded break at a desired site in the genome, followed by the insertion of the transgene via homology-directed repair (Wang et al., 2016). These synthetic biology tools for gene editing have significantly impacted the design of methods for transgene integration and cell line engineering as they eliminate the need to develop host cell lines containing the recombinase sites. CRISPR/Cas9-mediated site-specific integration was recently used to generate CHO cells containing the gene encoding the anti-PD1 monoclonal antibody integrated at a transcriptionally active site, reducing the timeline of cell line development and enhancing antibody titers (Zhao et al., 2018). One of the advantages of CRISPR-Cas9 over TALENs and ZFNs is its use of a single guide RNA, which significantly simplifies the experimental procedures. Tailoring TALEN and ZFN systems to a specific target requires re-engineering of the TALEN/ZFN protein while adapting CRISPR-Cas9 systems involves only the redesign of the 20-bp recognition sequence within the gRNA. Such procedures can be particularly involved for TALENs as they require the re-design of both the DNA-binding and nuclease domains.

By providing control over the site of integration and transgene copy number, methods for targeted transgene integration at specific genomic loci enable precise control over the expression level of recombinant proteins with minimal effects on cellular functions that are otherwise often observed upon disruption of chromosomal integrity and integration of foreign genetic material. The productivity of the resulting cell lines depends on the efficiency of screening and validation strategies. The main requirement for screening and validation strategies is the isolation of integration sites that allow stable integration of large sequences with high efficiency and that respond to orthogonal tools for control of gene expression. Leveraging modern genome editing tools for targeted transgene integration has thus the potential to dramatically improve the efficiency of integration of genetic sequences of desired size and the generation of stable cell lines for recombinant protein production. Programmable gene delivery technologies based on engineered Cas9-PiggyBac fusions combine the specificity of Cas9 with the payload transfer efficiency of the PiggyBac system to enable precise delivery of a wide range of payloads with minimal off-target events (Pallarès-Masmitjà et al., 2021). A similar strategy involving PRIME editing combined with site-specific recombinases (SSRs) was developed to achieve targeted integration of DNA plasmids of over 5000bp (Anzalone et al., 2022). A system based on Cas9 combined with recombinase-mediated cassette exchange (RCME) was used for multigene expression, including a therapeutic monoclonal antibody and unfolded protein response (UPR) genes expected to enhance recombinant protein production (Shin et al., 2021). This evidence points to the great, and possibly largely untapped potential of synthetic biology techniques for targeted gene integration for improving recombinant protein production by enabling the precise delivery of large genetic payloads.

2.2. Promoters, enhancers, and insulators

Transgene expression depends mainly on regulatory sequences controlling transcriptional processes, including promoter and enhancer sequences. High transgene expression is typically achieved using viral promoters, namely the human cytomegalovirus (CMV), Rous sarcoma virus (RSV), and Simian virus 40 (SV40) promoters. If the promoter size is a concern, the RSV and SV40 promoters, which are shorter in length, are typically preferred. Despite providing high transgene expression levels, the CMV promoter is prone to cell line-dependent methylation-induced gene silencing, presents cell-cycle dependency, and variable transgene expression levels (Bruening et al., 1998; Hsu et al., 2010; Kim et al., 2011). Long-term transgene expression is obtained using eukaryotic promoters such as the Chinese hamster elongation factor-1 alpha (CHEF-1α), human elongation factor-1 alpha (EF-1α), human ubiquitin C (UBC), β-globin, and phosphoglycerate kinase (PGK). The EF-1α and the cytomegalovirus early enhancer chicken/β-actin (CAG) promoters provide consistent transgene expression independent of cell type (Hitoshi et al., 1991; Qin et al., 2010). CAG is a synthetic promoter composed of the CMV enhancer and a rabbit β-globin intron with a chicken β-actin transcription start site that allows for high transgene expression (Hitoshi et al., 1991).

Inducible expression systems can be used to achieve control over target protein expression. Based on the tetracycline transcriptional activator (tTA), the Tet-OFF and Tet-ON systems are the most widely used inducible systems that enable user control through the addition of the small molecules tetracycline or doxycycline (Pedone et al., 2019). Activation of gene expression in these systems is mediated by binding of the small molecule inducer to the transcription factor, which results either in the displacement of the transcription factor from the operator and de-repression of gene expression, or binding of the transcription factor to the operator, recruitment of the polymerase and activation of gene expression. Leveraging the modularity of these systems, advances in transcription factor engineering have widened the array of small-molecule inducible systems available to the mammalian cell engineering community, providing appealing features such as orthogonality and tunability (Kallunki et al., 2019), which may improve the design of cell factories. Light-inducible promoters provide an alternative method to control gene expression with greater temporal resolution than small-molecule inducible promoters. A synthetic light-dependent transcription factor was generated using Vivid (VVD), a small protein with a light-oxygen-voltage (LOV) domain that dimerizes in response to blue light. The engineered tripartite fusion protein consisting of the DNA-binding domain of the Gal4 transcription factor, VVD, and the p65 activation domain-containing transcriptional activator GAVP (Wang et al., 2012) is activated upon cell exposure to blue light, resulting in the recruitment of the activator to the Gal4-responsive promoter. Similarly, the “LITE system” is a light-inducible transgene expression system that comprises a programmable TALE DNA-binding domain fused to light-sensitive protein cryptochrome 2, and the transcriptional activator VP64 fused to calcium and integrin-binding protein 1 (CIB1) (Konermann et al., 2013). Upon illumination with blue light, CRY2 associates with the CIB1-VP64 fusion protein, leading to the recruitment of the transcriptional machinery. Overall, these systems enable precise, reversible control over gene expression with increased temporal resolution compared to chemically inducible systems. An optogenetic OFF-switch was also developed for remote-controlled downregulation of protein expression (Baaske et al., 2018). A blue-light responsive optogenetic OFF-switch, Blue-OFF, was designed using a light-responsive repressor comprising a photosensitive transcription factor EL222 fused to transcriptional repressor KRAB. The reporter gene was placed under the control of a promoter followed by five copies of an EL222-binding domain. Upon exposure to blue light, EL222-KRAB dimerizes and represses the transcription of the reporter gene. This system enables fast and fully reversible repression of a target gene, expanding the synthetic biology toolkit for light-induced regulation of gene expression, which was previously focused only on ON switches. Overall, both chemical and light-inducible systems provide useful tools for controlling the temporal dynamics of gene expression and can be employed to control the production of recombinant proteins with greater temporal resolution.

Enhancers are cis-acting genetic elements that augment the promoter activity and boost the expression of the transgene. They typically range from 50 bp to 1000 bp in size and function independently of their position and orientation (Pennacchio et al., 2013). Enhancers contain binding sites for transcription factors and mediate gene expression by increasing the probability and/or rate of transcription initiation by opening the chromatin domain and facilitating the recruitment of the transcription machinery to the promoter (Raab and Kamakaka, 2010). Enhancers are usually separated from their promoters by thousands of base pairs and contain binding sites for transcription factors, which form clusters, thus augmenting the formation of a multiprotein complex that includes RNA polymerase and other proteins that make up the transcription machinery. This complex mediates the direct interaction of enhancer and promoter elements through enhancer- and promoter-bound factors, resulting in control of gene expression (Blackwood and Kadonaga, 1998). Engineering enhancer sequences offers new opportunities for the design of synthetic transcriptional regulatory sequences, as the activity of enhancers can be modulated by altering the number and specificity of the transcription factor binding sites (Schlabach et al., 2010). Designing regulatory sequences with the appropriate combination of enhancer and core promoter sequence provides an additional opportunity to achieve the desired level of transgene expression. The combination of the human cytomegalovirus immediate-early core promoter element (hPCE) with the EF-1α promoter, for instance, increased the GFP expression in CHO cells relative to the vector containing only the CMV core promoter (Wang et al., 2018).

Due to the DNA’s three-dimensional folding, enhancers may bypass neighboring genes and affect the regulation of genes located farther on the DNA sequence. Synthetic biology approaches have been recently explored to design strategies that circumvent this issue. Insulators prevent interference between neighboring sequences and protect promoters from interacting with enhancers outside of the insulator sequences. Insulators can be generally classified as enhancer-blockers, which block the communication between the enhancer and the promoter through the formation of chromatin loops, and barrier insulators, which prevent promoter silencing by disrupting heterochromatin formation⁴⁴. Incorporating insulators such as the chicken globin locus control region hyper-sensitive site-4 (cHS4) insulator into vectors for the expression of recombinant proteins ensure productive enhancer-promoter interactions and prevents epigenetic silencing, ultimately enhancing the stability of transgene expression (Maksimenko et al., 2015; Romanova and Noll, 2018; Takagi et al., 2017).

Temporal control over gene expression is typically achieved using inducible promoters. The recent development of a wide array of synthetic transcription factors has opened the way to new opportunities for the development of inducible expression systems that offer precise and tunable control of recombinant protein expression. Synthetic transcription factors are based on fusion proteins consisting of a DNA-binding domain and a regulatory domain. The DNA binding domain recognizes an operator sequence with user-defined specificity and affinity and the regulatory domain determines the function of the transcription factor as a transactivating or trans-silencing regulator (Ausländer and Fussenegger, 2013; Hörner and Weber, 2012). Most synthetic transcription factors are regulated by small molecules that alter the transcription factor binding affinity for the operator. Transgene expression can be controlled by altering the promoter design, and, specifically, the number of operator sites and spacing between the operator sites and the promoter and transcription start site (Garcia et al., 2012). Temporal control of transgene expression is also easily achieved by controlling the dosage of small-molecule inducers. Modulation of the transcription factor-operator interaction through appropriate design of the operator sequence or transcription factor engineering offers additional avenues to control transgene expression⁵ (Engstrom and Pfleger, 2017; Garcia et al., 2012; Liu et al., 2019). Tunable control of recombinant protein expression is a particularly desirable feature for large-scale cultures, as it allows separating the cell growth phase from the protein production phase, which is considered an ideal approach to maximizing production.

2.3. Other regulatory elements that enhance the expression of recombinant proteins

Because the location of a chromosomal locus is known to influence gene activity, negative positional effects are expected to affect transgene expression over time. To overcome gene silencing effects and the resulting reduction in protein production, integration cassettes are designed to include regulatory elements that provide epigenetic control by reducing the spread of heterochromatin states into regions of euchromatin or by rendering chromatin transcriptionally permissive (Richards and Elgin, 2002). These elements enhance promoter activity and protect the promoter from epigenetic silencing, thus maintaining sustained transcription levels over prolonged culturing periods.

Scaffold or matrix attachment regions (S/MARs) are epigenetic regulatory elements used to achieve consistent transgene expression. They consist of AT-rich DNA sequences that bind to the nuclear matrix, resulting in the formation of chromatin loops, thus modifying the chromatin architecture (Heng et al., 2004). S/MAR sequences range from 300 bp to 1.5 kb in length and are placed upstream of the promoter controlling the expression of the transgene. By recruiting transcription factors and chromatin remodeling enzymes, they mediate control of transcription and insulation of the transgene from epigenetic silencing (Harraghy et al., 2008). The addition of the human β-globin MAR and β-interferon MAR sequences to an expression vector regulated by the SV40 promoter, for instance, was found to enhance transgene expression stability over a prolonged culturing period. S/MAR elements also increased the transfection efficiency, resulting in an increased fraction of the cell population presenting stable transgene expression (Saunders et al., 2015; Zhao et al., 2017). The efficiency of S/MARs, however, is likely cell line-dependent, pointing to the need for testing combinations of promoters and S/MAR elements for case-by-case optimization of S/MARs to regulate recombinant protein production in the desired host cell line.

Stabilizing anti-repressor (STAR) elements are regulatory sequences ranging from 500 bp to 2 kb in length that flank the transgene and stabilize the transgene expression by counteracting the effect of chromatin-associated repressors. STAR elements STAR7 and STAR40 have been used to enhance transgene expression in the context of a screen for the selection of cells engineered to express antibiotic resistance selection genes in which STAR elements were found to prevent gene silencing and increase antibiotic resistance (Kwaks et al., 2003; Saunders et al., 2015).

Ubiquitously acting chromatin opening elements (UCOEs) are methylation-free sequences that maintain the chromatin in a transcriptionally active state independently of the site of integration. They have been included in the design of cell lines engineered to prevent the spread of chromatin methylation and deacetylation at desired genetic sequences and limit transgene silencing. Specifically, the integration of A2UCOE improved the production of the anti-CD20⁺ antibody in CHO cells 6 folds and resulted in sustained antibody production over prolonged periods (Rocha-Pizaña et al., 2017). A direct comparison of the insulator (cHS4), S/MAR (MARX_S29), STAR (STAR40), and UCOE (A2UCOE) chromatin-modifying elements within the same vector context revealed that the UCOE (A2UCOE) element to be the most efficient in enhancing the level of transgene expression and stability of transcription, leading to a 6.75-fold increase in antibody production in CHO cells (Saunders et al., 2015).

The use of multiple regulatory elements that enhance the efficiency of transcription and prevent transgene silencing results in the design of large expression vectors, often affecting the efficiency of transfection or exceeding the packaging capacity of viral vectors. Altering the epigenetic environment of the chromatin surrounding the site of integration provides an alternative approach to preventing epigenetic silencing of the transgene. This approach involves the use of a fusion protein consisting of a moiety mediating chromatin modification and a moiety mediating targeting of the fusion protein to the desired chromosomal site. Specifically, the expression of a fusion protein consisting of the p300 histone acetyltransferase (HAT) domain and the Lex-A protein was found to mediate the recruitment of the p300 HAT domain to the CMV promoter containing Lex-A binding sites and, as a result, to enhance transgene expression (Kwaks et al., 2005). Most strategies to enhance transgene expression provide significant but often limited results: leveraging different mechanisms by combining multiple regulatory elements typically provides additive or even synergistic results. A significant increase in transgene production stability, for instance, was obtained by incorporating the anti-repressor elements in the expression vector containing the Lex-A binding sites that recruit the p300 HAT domain to the CMV promoter (Kwaks et al., 2005). Generally speaking, multiple chromatin-modifying elements, when used in association with strong promoters, prevent DNA methylation-induced silencing and provide a stable transgene expression (Kwaks et al., 2005).

Tools that operate at the post-transcriptional level also affect the transgene expression by enhancing the stability and translation of primary transcripts. The efficiency of mRNA processing can be enhanced using post-transcriptional regulatory elements (PTREs). The use of PTREs such as a tripartite leader sequence of the 5’ untranslated region (UTR) of human adenovirus mRNA and Woodchuck hepatitis virus post-transcriptional regulatory element (WPRE) under the control of a CMV promoter resulted in a ~9-fold increase in IFN-γ and a ~5-fold increase in trastuzumab expression in HEK 293 cells (Carswell and Alwine, 1989; Mariati et al., 2012; Miller, W. L., Martial, J. A., Baxter, J.D., 1985). Other PTREs have been explored to enhance mRNA stability and translation and include polyadenylation signals, 5’ UTR of the human heat shock protein 70 mRNA (Hsp70), and introns (Mariati et al., 2012).

2.4. Regulatory elements that control the expression of proteins with quaternary structure

The production of complex proteins such as monoclonal antibodies often involves the co-expression and assembly of multiple protein subunits. The expression of quaternary proteins requires co-transfection of multiple vectors or single vectors for the expression of multiple transgenes under the control of different promoters (e.g., the light chain [LC] and heavy chain [HC] of monoclonal antibodies). In the multi-promoter single vector design, each transgene is transcribed independently; as a result, this method lacks control over the relative expression levels of LC and HC (Rita Costa et al., 2010; Wurm, 2004). The antibody titers and quality were found to vary greatly depending on the LC-to-HC expression ratio (Chusainow et al., 2009). This observation points to the need to control the relative expression levels of protein subunits for maximizing the production of complex recombinant proteins with quaternary structures.

Strategies to link the expression of multiple polypeptide chains include methods to express fusion proteins that are subsequently processed to release independent subunits, as well as methods for producing polycistronic mRNAs that are translated into multiple subunits. The first strategy is typically implemented by creating polypeptide chains fused using 2A peptides: translation of one open reading frame followed by self-cleavage of the 2A peptide results in the release of equal amounts of the protein subunits. Self-cleavage of the 2A peptide leaves a two-amino acid scar at the site of cleavage that in most cases does not affect the protein product. The two-amino acid scar can be eliminated through the addition of a furin recognition sequence that mediates the removal of the 2A residues from the final protein product (Shaimardanova et al., 2019). Self-cleavage of a 2A peptide results in the formation of LC and HC and native processing and assembly of full-length antibodies. Overexpression of LC and HC genes linked by 2A peptides, however, is plagued by some degree of inefficient 2A peptide cleavage, which may result in the accumulation of HC-2A-LC or LC-2A-HC fusion proteins and the formation of insoluble aggregates of the resulting off-pathway misfolded intermediates. The need to optimize the expression of properly processed LC and HC has motivated the characterization of several 2A peptide variants that present sequence-dependent cleavage efficiency. Interestingly, a strong correlation between the cleavage efficiency of the 2A peptides and the resulting antibody yields was observed (Chng et al., 2015; Liu et al., 2017).

Methods for producing constant ratios of multiple protein subunits are also based on the use of internal ribosome entry sites (IRES), which offer an appealing alternative to 2A peptides. IRES introduces a ribosome binding site within an mRNA sequence, resulting in an additional site for initiation of translation, which proceeds in a cap-independent manner, i.e., without requiring a 5’ cap for the translation initiation factors. Most IRES, however, result in lower expression of the gene 3’ of the IRES (Mizuguchi et al., 2000). Because an excess of LC levels was found to be required for maximal yields of recombinant antibodies and reduced aggregation of off-pathway intermediates, strategies based on LC-IRES-HC expression systems may provide a solution to the optimization of antibody production yields (Ho et al., 2013; Wickramasinghe and Laskey, 2015).

In an attempt to achieve controlled expression of LC and HC with excess levels of LC compared to HC, tricistronic expression systems, consisting of LC and HC encoding genes and an antibiotic selection cassette separated by two different IRES were also explored (Balasubramanian et al., 2016; Ho et al., 2012). Such expression systems designed to produce the LC from the first cistron and the HC from a second cistron ensure optimal ratios of expression of LC and HC and minimal aggregate formation, thereby enhancing antibody titers. Placing the selection cassette downstream of an IRES results in the expression of the upstream gene at higher levels than the selection marker required for cell survival, ultimately leading to the selection of high-producing cells. Attempts to optimize the production of recombinant proteins have also involved exploring alterations in the IRES sequence, which allow for modulating the expression ratios of the LC, HC, and selection marker (Chai et al., 2018; Sadikoglou et al., 2014; Yeo et al., 2017). Overall, compared to 2A peptides, IRES-based expression systems for the production of monoclonal antibodies typically result in reduced aggregation and higher yields of high-quality product (Ebadat et al., 2017).

2.5. Engineering translation and trafficking

The expression of recombinant proteins is often plagued by translational silencing due to the phosphorylation of the eukaryotic translation initiation factor 2 (eIF2) that typically occurs upon transfection and transduction procedures (Xiao et al., 2014). Cell engineering with exogenous plasmids also results in the activation of the translation inhibitor double-stranded-RNA-dependent protein kinase (PKR). Expression of the Ebola virus protein 35, which blocks activation of PKR was found to prevent translational silencing, specifically resulting in a 10-fold increase in the expression of recombinant protein TPL-2 complex in HEK 293 cells (Gantke et al., 2013). Similarly, overexpression of the eukaryotic initiation factor eIF3i subunit in HEK 293 cells was observed to cause a substantial increase in luciferase expression (Zhang et al., 2007). Such result was attributed to eukaryotic initiation factor (eIF3)-mediated upregulation in translation initiation and elongation which is likely to enhance global protein synthesis rates as well as cell proliferation (Roobol et al., 2020), but was also found to affect recombinant protein production in a cell type-specific fashion (Jossé et al., 2016; Mead et al., 2012).

The overexpression of recombinant proteins can cause a considerable burden on protein translation. Such a burden is expected to cause the activation of a global stress response that manifests through the activation of a series of signaling pathways to prevent cytotoxicity, including silencing of the transgene expression and reduction in global protein synthesis, as mentioned above. A way of circumventing translational burden while increasing recombinant protein production is to use non-coding RNAs to tune the translation of the target mRNA (Vito and Smales, 2018). Specifically, microRNAs (miRNAs) are transcribed as long primary transcripts but processed to yield 20–23 nucleotide non-coding RNAs that interact with the 3’ UTR region of the target mRNA, inhibiting translation or initiating degradation. A single miRNA can target multiple mRNAs via imperfect base pairing and hence modulate multiple mRNAs and cellular pathways. While the mechanisms of action are unclear, overexpression of mi-RNAs has been shown to increase recombinant protein production. For instance, expression of miR-557 and miR-143 resulted in a 2-fold and 1.2-fold increase in antibody production, respectively (Fischer et al., 2017; Schoellhorn et al., 2017). A potential explanation for this mi-RNA-mediated effect on recombinant protein production is mi-RNA-mediated targeting and downregulation of genes involved in apoptosis (Inwood et al., 2018). Another strategy to improve recombinant protein production leveraging RNA regulation consists in engineering long non-coding RNAs (lncRNAs, more than 200 nucleotides in length) that upregulate transcription in a gene-specific manner and promote translation of partially overlapping sense coding mRNA using an inverted SINEB2 sequence. Synthetic SINEUPs contain a 5’ DNA binding domain overlapping with the protein-coding target and the embedded inverted SINEB2 element that serves as the effector domain and functions as a translation activator (Carrieri et al., 2012; Zucchelli et al., 2016). The modularity of the synthetic SINEUPs allows for designing leader-specific binding domains to upregulate secreted proteins to produce multiple recombinant proteins in parallel. This approach has previously resulted in a 2–5-fold increase in the expression of recombinant proteins including antibodies (Carrieri et al., 2012; Sasso et al., 2018).

Most therapeutic proteins such as antibodies are extracellular proteins that are normally processed through the secretory pathway, beginning with co-translational translocation into the endoplasmic reticulum (ER), trafficking through the Golgi apparatus where glycosylation takes place, and culminating in secretion. Translocation into the ER lumen is mediated by 5 to 30-amino acid long signal peptides and typically represents one of the rate-limiting steps in the overall protein production process, explaining the observed lack of correlation between mRNA transcript levels and the yields of functional recombinant proteins. The signal peptide of nascent polypeptide chains is recognized by the signal recognition particle (SRP), resulting in the formation of an SRP-ribosome-nascent chain complex. The SRP mediates translocation of the complex into the ER, which is followed by cleavage of the signal peptide (Owji et al., 2018). Overexpression of recombinant proteins is typically achieved through protein fusion to well-characterized signal peptides known to result in high secretion efficiency. Miscleavage of the signal peptide has been observed in a significant fraction of protein chains and results in the truncation or elongation of the HC or LC, causing misfolding and accumulation of heterogeneous products, and ultimately lowering the overall product quality (Gibson et al., 2017). Signal peptides have been engineered to improve the affinity to the SRP, which correlates with the translocation efficiency and yield of the recombinant protein. The azurocidin signal peptide was found to increase IL-21 secretion approximately 2.5-fold and the Igκ signal peptide to enhance the expression of recombinant coagulation factor VII 1.5-fold in CHO cells (Cho et al., 2019; Peng et al., 2016). Similarly, optimization of the signal peptide mediating secretion of antibodies in CHO cells resulted in an increase in the yields of anti-HER2 antibody (Herceptin, 2.2-fold), anti-CD20 antibody (Rituxan, 2-fold), and anti-VEGF-A antibody (Avastin, 3-fold) (Haryadi et al., 2015; You et al., 2018). Unfortunately, the yield of antibody production was found to vary not only with the specificity of the signal peptide sequence but also with the specific type of antibody and the host cell line (Kober et al., 2013; Ramezani et al., 2017). As a result, the efficiency of translocation of signal peptides is currently tested and optimized on a case-by-case basis.

2.6. Recapitulating native post-translational modifications

The nature of a protein’s glycosylation patterns may affect its stability, binding interactions, and potentially even activity. In addition, glycosylation affects the pharmacokinetics of therapeutic proteins in blood circulation. The choice of host cell type for biomanufacturing involves considerations related to the need to recapitulate native glycosylation reactions as well as avoid glycosylation patterns that may elicit an immune response in humans. CHO cells, for instance, generate N-linked glycans that are recognized as foreign by the human immune system and that elicit an immune response to therapeutic proteins. Similarly, murine and other hamster-derived cells produce glycans with galactose-α1,3-galactose (α-gal) and N-glycolylneuraminic acid (Neu5Gc) terminals, which are also immunogenic in humans (Lalonde and Durocher, 2017).

Glycosylation patterns of protein therapeutics can be modulated by controlling the culture conditions. For instance, dimethyl sulfoxide (DMSO) and glycerol have a stabilization effect on recombinant proteins; chemical chaperones also stabilize proteins and prevent aggregation due to thermal stress, and low temperatures may reduce growth rate but enhance specific protein productivity (Rodriguez et al., 2008). Synthetic biology methods represent another strategy for glycoengineering, and compared to traditional glycoengineering methods have the potential to generate more specific and complex glycosylation modifications (Kightlinger et al., 2020). Because glycosylation affects antibodies’ function, altering the glycosylation pattern of monoclonal antibodies may be useful for therapeutic applications. The effector functions of antibodies include antibody-dependent cellular cytotoxicity (ADCC) and complement-dependent cytotoxicity (CDC). The ADCC activity is mediated by the binding of the antibody constant region (Fc) to the lymphocyte receptors (FcγRs) and is influenced by the glycan structure on the Fc region. The binding affinity of the antibody’s Fc region to the FcγRIIIa receptor of the natural killer cells increases in the absence of Fc fucosylation, resulting in enhanced ADCC. Similarly, the antibody’s sialylation profile affects its half-life in circulation (Bas et al., 2019), the mannosylation patterns influence its pharmacokinetics (Liu, 2015), and the galactosylation regulates CDC (Peschke et al., 2017). Reducing the fucose residues on the Fc region of recombinantly produced antibodies is expected to trigger ADCC and is thus considered a promising strategy to increase the efficacy of therapeutic antibodies. Knocking out or downregulating the FUT8 gene reduces antibody fucosylation. Specifically, the siRNA-mediated knockdown of FUT8 resulted in an 80% reduction in FUT8 mRNA and a 100-fold increase in the ADCC of the anti-CCR4 antibody (Mori et al., 2004). Other approaches have been explored to reduce fucosylation by decreasing the expression of FUT8 mRNA and include the expression of β1,4-mannosyl-glycoprotein 4-β-N-acetylglucosaminyltransferase III (GnTIII) and Golgi mannosidase II (ManII) or the bacterial enzyme GDP-4-dehydro-6-deoxy-D-mannose reductase (RMD) (Ferrara et al., 2006; von Horsten et al., 2010; Roy et al., 2018). Alternatively, the cellular fucosylation activity was completely removed by disrupting the fucosyltransferase through FUT8 knockout, which was achieved using CRISPR/Cas9 and ZFN genome editing methods (Malphettes et al., 2010; Ronda et al., 2014).

The addition of sialic acid residues to monoclonal antibodies endows their anti-inflammatory properties and improves their residence time in circulation by preventing recognition by the asialyl glycoprotein receptors in the liver (Durocher and Butler, 2009). Sialylation of the recombinant therapeutic can be increased either by knocking down the expression of sialidases (e.g., Neu2, Neu 3) or overexpressing glycosyltransferases (e.g., ST3GALI, ST3GALIV, and ST6GAL1) that mediate sialylation reactions (Zhang et al., 2010a; Zhang et al., 2010b). Specifically, the overexpression of ST6GAL1 resulted in an increase in the sialylation profile of the anti-EGFR × anti-CD bispecific antibody in CHO cells (Onitsuka et al., 2012). Overexpression of the glycosyltransferase β−1,4-galactosyltransferase (β4GALT1) was found to enhance the galactosylation of recombinant proteins, which is known to affect the CDC of therapeutic antibodies. Transient concomitant expression of ST6GAL1 and β4GALT1 in CHO cells enhanced the sialylation and galactosylation of trastuzumab in CHO cells (Raymond et al., 2015). In a recent study, host cells were engineered for user-controlled modulation of the glycosylation and fucosylation of an IgG1antibody by first generating a ST6GAL1 and β4GALT1 knock-out CHO cell line and then integrating genetic circuits expressing synthetic glycosyltransferase genes under the control of small molecule-inducible promoters (Chang et al., 2019).

The loss of patent protection for previously developed therapeutic antibodies enables the development of biosimilars, a field in which glycoengineering methods can be employed to reproduce the properties of the original molecules to create less expensive drugs (Duivelshof et al., 2019). Synthetic biology approaches for glycoengineering allow enhancing different aspects of therapeutic proteins, such as productivity and stability. For instance, levels of specific glucosyltransferases (GTs) and sugar donor metabolism-related enzymes can be modulated using gene editing technologies resulting in the production of more homogeneous glycosylation patterns in the final products, thereby increasing the production yields (Kightlinger et al., 2020). Ultimately, these novel approaches offer the opportunity to improve the yield and quality of therapeutic products, thus providing more effective and accessible treatments.

3. CELL ENGINEERING TO INCREASE THE PRODUCTION OF RECOMBINANT PROTEINS

3.1. Controlling cell proliferation

Because cells cycle through different phases characterized by different cell growth and metabolic activity, regulating the cell cycle allows controlling the production of recombinant proteins (Figure 3). Specifically, a widely explored strategy consists in regulating the cell progression from a state of proliferation to maximize viable biomass, arresting the cell cycle to prioritize productivity. Such an approach capitalizes on methods to prevent cell division through cell cycle arrest and thus without inducing cell death and affecting protein synthesis. As cell cycle progression is arrested in G1, cells present an increase in metabolic activity as well as an increased size and expression of genes responsible for ribosome biosynthesis. Cyclin cycle division 25 (CDC25) phosphatases, specifically CDC25 homolog A (CDC25A), activate cyclins and cyclin-dependent kinases (CDKs), which, in turn, regulate cell cycle transitions and are specifically involved in G1-to-S and G2-to-M transitions. Overexpression of CDC25A (Lee et al., 2013) enables cells to bypass cell cycle checkpoints and promotes cell cycle progression. Overexpression of CDC25A resulted in a 2.4-fold increase in the specific productivity of the anti-EGFR x anti-CD3 bispecific single-chain diabody in CHO cells. Similarly, conditional expression of the CDC25B (Matsuyama et al., 2015) using the Cre/loxP recombination system resulted in a 15-fold higher specific productivity of bispecific antibody scDb-Fc in CHO cells. A potential pitfall of this strategy is that cells engineered with CDC25A and CDC25B bypass DNA damage-induced checkpoint arrest, which may lead to increased apoptosis through genomic instability.

Figure 3. Cell engineering approaches to improve biomanufacturing:

(1) Controlling cell proliferation; (2) Engineering survival pathways; and (3) Engineering protein secretory pathways.

Cyclin-dependent kinases inhibitors (CDKIs) inhibit the G1-to-S phase transition, resulting in the arrest of the cells in G1. Expression of CDKIs, such as p21^Cip1 and p27^Kip1, which increases as a result of histone deacetylation, inhibits the downstream phosphorylation of retinoblastoma protein (Rb), a tumor-repressing protein, resulting in G1 cell cycle arrest (Sunley and Butler, 2010). Transient overexpression of p21^Cip1 and p27^Kip1 in CHO cells resulted in a 5-fold and 4-fold increase in SEAP production (Fussenegger et al., 1997), respectively. These methods lead to an increase in overall protein productivity since cells in growth arrest do not employ cellular resources for biomass production. However, a rresting cell growth in G1 causes the activation of complex signaling pathways leading to cellular states that may affect the product quality and could prevent commercial applications based on these techniques (Jazayeri et al., 2018).

An alternative engineering strategy aimed at improving recombinant protein production by increasing cell proliferation is based on the manipulation of mTOR, a master metabolic regulator that plays a significant role in cell growth and proliferation. Overexpression of mTOR (Dreesen and Fussenegger, 2011) in CHO cells led to an increase in cell size, proliferation, viability, and specific productivity, resulting in a considerable increase in therapeutic antibody titers. Another approach to controlling cell growth for enhancing recombinant protein production is based on the overexpression of miRNAs involved in the regulation of cellular proliferation. Expression of miR-30 and miR-17 (Jadhav et al., 2012), two microRNAs associated with enhanced cell proliferation, led to an increase in cell density and an almost 2-fold increase in the titer of erythropoietin-Fc in CHO cells. A potential issue associated with strategies aimed at increasing cell proliferation is cell growth saturation, which leads to nutrient depletion, hypoxia, and accumulation of waste products, the primary cause of cell death through apoptosis in large-scale cultures.

3.2. Engineering survival pathways

Culturing cells under conditions that maximize the production of recombinant proteins typically culminates in activations of pro-apoptotic triggers, namely nutrient deprivation, hypoxia, waste product accumulation, and mechanical agitation. A reduction in the number of viable cells inevitably affects the yields of recombinant therapeutic proteins. In addition to strategies for controlling cell growth rate through bioreactor design and modulation of culturing conditions, cell engineering approaches for delaying apoptosis and prolonging protein production have been explored and include mainly attempts to regulate the expression of gene encoding pro- or anti-apoptotic proteins or chemical modulation of key regulatory proteins such as caspases (Figure 3). Similar to chaperone engineering, the overexpression of anti-apoptotic genes including BCL2, MCL1 (Majors et al., 2009), BCL2L1, AVEN, XIAP, and CRMA has resulted in protein-specific results and needs to be tested on a case-by-case basis. The transient overexpression of BCL2L1 or MCL1 resulted in a 1.82-fold and 1.5-fold increase in the titers of anti-PD1 (Zhang et al., 2018) antibodies in CHO cells. The downregulation of pro-apoptotic genes, namely BAX and BAK, is typically achieved using small-interfering RNA (siRNA) or gene knockout. BAX and BAK knockout cells generated with ZFNs displayed enhanced cell survival and a 35% increase in IFN-γ production compared to parental CHO cells (Cost et al., 2010). Similarly, downregulation of pro-apoptotic genes Caspase-3 and Caspase-7 using siRNA increased cell viability and culture longevity of CHO cells and resulted in a significant increase in thrombopoieti n (Sung et al., 2007) production. Inhibition of the pro-apoptotic miRNA Mmu-miR-466h-5p (Jazayeri et al., 2018) also increased recombinant protein production. Taken together, these strategies suggest that apoptotic pathways play a key role in the fitness of cells engineered for high yield production of recombinant proteins, but more sophisticated systems-level approaches are required to generate universal strategies for modulating apoptosis that would overcome current limitations associated with protein- and cell type-specific variability.

3.3. Engineering the protein secretory pathway

Post-translational modifications and processing of newly synthesized proteins as they are trafficked through the compartments of the secretory pathway present a bottleneck in protein production. Proteins destined for extra-cytoplasmic compartments contain an N-terminal signal sequence that guides newly synthesized polypeptide chains to the endoplasmic reticulum (ER). After SRP-mediated translocation from the cytoplasm to the ER, proteins are processed through a set of folding and post-translational modifications, such as glycosylation. Secretory proteins are released from the ER upon packaging into vesicles that mediate transport to the membrane compartments of the Golgi apparatus and subsequently to the plasma membrane for secretion to the extracellular space through exocytosis.

The translocation of proteins through the secretory pathway is regulated through a series of checkpoints that maintain quality control. Within the ER, chaperones mediate quality control by ensuring the folding of proteins into their native three-dimensional structures. Proteomic analyses of cells engineered to overexpress recombinant proteins showed elevated levels of the main ER chaperones, namely binding immunoglobulin protein (BIP), glucose-regulated protein-94 (GRP94), and protein disulfide isomerase (PDI) (Smales et al., 2004). However, overexpressing these ER chaperones has not resulted in a consistent increase in the yield of recombinant proteins. For instance, Increasing BIP levels through gene overexpression does not affect the production yields (Smales et al., 2004). Overexpressing PDI in cells engineered to express different model proteins has led to protein-specific results; interestingly, it decreased the secretion of TNFR:Fc and did not affect that of IL-15 (Davis et al., 2000; Mohan et al., 2007). Co-expressing PDI and the PDI isoform Erp57 resulted in a 2.1-fold increase in the accumulation of thrombopoietin in CHO cell (Mohan et al., 2007). Simultaneous overexpression of multiple proteins of the heat shock family (e.g., Hsp27 and Hsp70) increased the yield of IFN-y production by 2.8-fold (Lee et al., 2009). Overexpressing a single ER chaperone, even when successful in increasing the yields of recombinant protein production, may ultimately provide an inefficient strategy to meet the production needs as it may result in insufficient levels of the chaperone’s essential co-factors. Moreover, the overexpression of a single or even multiple chaperones may alleviate the pressure on the ER folding capacity but eventually creates a new rate-limiting step in the protein folding process, ultimately providing limited enhancement of the recombinant protein yields, as observed upon co-expression of the main ER chaperones PDI and ERO1 in antibody-expressing CHO cells (Mohan and Lee, 2010). Strategies based on the simultaneous overexpression of multiple chaperones may provide an interesting avenue for protein expression but are also likely to require more sophisticated synthetic biology approaches to engineer designer cells that sense the folding needs and respond by enhancing the cellular folding capacity at specific steps of the secretory pathway.

The aberrant accumulation of misfolded protein in the ER results in proteotoxic stress and triggers activation of the unfolded protein response (UPR), a coordinated series of transcriptional and post-translational pathways aimed at reducing the load of misfolded and unfolded proteins mainly through upregulation of chaperones and other cellular folding mechanisms and reduction of global protein synthesis to alleviate cellular burden (Hetz, 2012). If sustained, the UPR culminates in the activation of pro-apoptotic pathways. The accumulation of misfolded proteins in the ER leads to the activation of three ER transmembrane proteins, namely protein kinase R (PKR)-like endoplasmic reticulum kinase (PERK), activating transcription factor 6 (ATF6) and inositol-requiring enzyme 1 (IRE1), which function as sensors of proteotoxic stress. Under resting conditions, the sensors are bound to the molecular chaperone-binding immunoglobulin protein (BiP). An increase in misfolded proteins in the ER lumen effectively results in the sequestration of BiP and release of the UPR sensors. The activation of PERK and IRE1 leads to the induction of signaling pathways aimed at restoring ER homeostasis. PERK is activated upon dimerization and autophosphorylation and results in the phosphorylation of the eukaryotic translation initiator factor 2α (eIF2α), which, in turn, mediates the translation of activating transcription factor 4 (ATF4), a transcription factor responsible for the transcription of genes involved in autophagy, apoptosis, and amino acid metabolism. IRE1 activation also proceeds through dimerization and autophosphorylation, which trigger its RNase activity and processing of the mRNA encoding X-box binding protein 1 (XBP1), a global regulator for sustained stress recovery across the endomembrane and endocytic systems and a general organelle biogenesis factor (Figure 3). Numerous strategies aimed at recapitulating the innate function of the UPR as a mechanism to alleviate cellular stress and restore homeostasis have been explored in an attempt to enhance the production of natively folded recombinant proteins. The ectopic expression of XBP1 in CHO-K1 cells (Tigges and Fussenegger, 2006) was found to induce a global increase in the cellular secretion capacity and, specifically, a 6-fold increase in production of the model protein SEAP, by induction of ER-localized DnaJ 4 (Erdj4), which leads to enlargement of the ER and Golgi organelles and increased protein secretion. Attempts to leverage XBP1 function and increase XBP1 expression to enhance the productivity of other recombinant proteins, namely monoclonal antibodies, interferon-γ (IFN-γ), and erythropoietin (EPO), in CHO-K1 cells (Ku et al., 2008), revealed that this strategy proves successful only when the host cell has reached a secretory bottleneck. In CHO-K1 cells with high concentrations of EPO-encoding plasmid, the overexpression of XBP1 resulted in up to a 2.5-fold increase in EPO titers. Taken together, these results suggest that the extent to which the overexpression of XBP1 affects the yields of recombinant protein production depends on the expression levels of the recombinant protein.

Downstream of the PERK signaling pathway, ATF4 controls the expression of UPR target genes, including GADD34 and CHOP. Overexpression of ATF4 significantly increased the production of recombinant antithrombin III (AT-III) in CHO cells, most likely by enhancing dephosphorylation of eIF2a, which releases translation attenuation caused by ER stress, through GADD34 (Ohya et al., 2008). Overexpressed GADD34 was found to increase the production of AT-III in CHO cells (Omasa et al., 2008).

Natively folded secretory proteins are sequestered into vesicles that regulate transport from the ER, through the Golgi, to the plasma membrane. Vesicular trafficking requires the superfamily of SNARE proteins, which mediate membrane fusion by anchoring both transport vesicles and target membranes. Vesicle fusion is also catalyzed by the Sec1/Munc18 (SM) family proteins. The ectopic expression of Sly1 and Munc18c, SM proteins involved in modulating the fusion of COPII vesicles with the Golgi apparatus and that of Golgi-derived secretory vesicles with the plasma membrane, was shown to increase the production of SEAP, SAMY, VEGF₁₂₁ and Rituximab in CHO cells (Peng and Fussenegger, 2009). Similarly, the ectopic expression of exocytic SNARE proteins SNAP-23 and VAMP8 was found to produce an increase in recombinant protein production in HEK 293, HeLa, and CHO-K1 cells, likely due to a global increase in the cellular secretion capacity (Peng et al., 2011). Another key component of secretory transport is the ceramide transfer protein (CERT), which mediates ATP-dependent ceramide transport from the ER to the Golgi complex and is a substrate of protein kinase D (PKD), a component of the trans-Golgi network (TGN)-to-plasma membrane transport system. Heterologous expression of CERT in CHO cells was found to increase exocytosis of single-chain protein HSA, monoclonal IgG1 subtype antibody (Florin et al., 2009), and tissue plasminogen activator (t-PA) (Rahimpour, 2013). While the de-regulated modulation of individual components of the secretory pathway has yielded improvements in the production of biopharmaceuticals, a synthetic biology approach for building dynamic control systems that regulate the expression of master regulators or key components of the secretory pathway may provide a more efficient strategy to maximize protein secretion, particularly through specially designed sensors for monitoring the activity of the secretory pathway and genetic control systems for dynamically tuning the secretion capacity to the cellular needs.

4. CONCLUSIONS

The biomanufacturing of therapeutic proteins is a complex process that requires considerable investment in time and resources to ensure high-production titers. The constant and increasing demand for therapeutic proteins points to the pressing need to improve cell factory engineering methods to overcome biomanufacturing limitations. As described in this review, the synthetic biology community has recently generated a wide range of tools that are poised to improve cell factory engineering at different operational levels of the biomanufacturing process. CRISPR-Cas9-based methods have tremendously improved the efficiency and specificity of transgene delivery. Libraries of genetic parts for transcriptional regulation, including promoters and enhancers, enable fine-tuning of transgene expression with great precision. Epigenetic regulatory elements such as S/MARs allow for minimizing epigenetic silencing, ultimately contributing to maintaining the desired expression levels over time. Engineering the regulatory mechanisms controlling translation and intracellular trafficking holds great potential to improve the yields of folded, active products, which is often the rate-limiting step of biomanufacturing. Finally, glycoengineering approaches provide a unique opportunity to ensure the production of stable proteins with innate immunogenicity and enhanced efficiency.

In addition to refining the toolkit of genetic parts for recombinant protein expression, efficient and sustained protein production requires manipulating complex cellular pathways involved in promoting protein post-translational processing and maintaining cell viability. Progress in our fundamental understanding of the regulatory mechanisms controlling homeostatic cellular processes including cell division, cell death, and protein trafficking has enabled a number of cell engineering approaches such as arresting the cell cycle in a state of high metabolic activity (i.e., G1), blocking cellular death to increase the protein production span and enhancing the protein trafficking capacity. These methods are currently mostly based on the deregulated modulation of key cellular components and typically culminate in bottlenecks in protein production and cellular stress. The design of dynamic cell factories engineered through the incorporation of orthogonal genetic circuits that sense relevant cellular processes and, in response, modulate recombinant protein production is expected to provide a more efficient method to improve biomanufacturing. Such an approach would enable limiting protein production to permissive cellular states, such as states of high metabolic activity, preventing cellular stress and, ultimately, generating cell factories that sustain the production of recombinant proteins.

In summary, the advances in synthetic biology combined with progress in our fundamental understanding of regulatory mechanisms that affect metabolic states and cell viability in cells engineered to produce therapeutic proteins will offer tremendous opportunities to innovate the design of cell factories that are able to cope with the production of complex therapeutic molecules as well as supramolecular complexes such as adeno-associated viruses.