Genetic optimization of recombinant glycoprotein production by mammalian cells

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Genetic optimization of recombinant glycoprotein production by mammalian cells Martin Fussenegger, James E. Bailey, Hansjörg Hauser and Peter P. Mueller Genetically modified mammalian cells are the preferred system for the production of recombinant therapeutic glycoproteins. Other applications include engineering of cell lines for drug screening and cell-based therapies, and the construction of recombinant viruses for gene therapy. This article highlights contemporary core genetic technologies and emerging strategies for genetically engineering mammalian cells for optimal recombinant-protein expression.

he production of recombinant secretory proteins in mammalian cells is affected by the growth media, culture conditions and a large variety of genetic factors1. The first step of gene expression, transcription, has been one of the main focuses in biotechnological applications using mammalian cells. These efforts resulted in efficient and regulated promoter– transactivator systems. The influence of the surrounding chromosomal DNA on promoter activities has only recently been addressed systematically using virusbased plasmids, transient expression and site-directed recombination and integration. Variations in the expression levels of the same construct in individual clones initiated the development of sophisticated, multicistronic expression vectors and screening procedures. In addition to the productivity of individual cells, the final production level depends on the cell number and on the productive lifespan of the culture, items addressed by the genetic engineering of cell growth and the suppression of programmed cell death. Apart from achieving high expression levels, product quality is of paramount importance. Post-translational modifications can impose significant problems for in vivo applications of the product. A major objective of current research is the use of metabolic engineering to achieve product protein modifications that are compatible with pharmaceutical applications.

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High-level, regulated gene expression Most current applications make use of cell lines with stably integrated genes under the control of a strong cellular or viral promoter2. Some cellular promoters (e.g. EF1a) or viral promoters, including the cytomegalovirus promoter3, can be used in different cell lines for high-level protein expression, but their transcriptional activity varies depending on the cellular levels of the relevant transcription factors and on the chromatin structure at the integration site. In addition, scaffold- or matrix-attachment regions (S/MAR elements) of chromosomal DNA can augment the M. Fussenegger and J. E. Bailey are at the Institute of Biotechnology, ETH Hönggerberg, HPT, CH-8093 Zurich, Switzerland. H. Hauser and P. P. Mueller ([email protected]) are at the Department of Gene Regulation and Differentiation, GBF – National Research Center for Biotechnology, Mascheroder Weg 1, D-38124 Braunschweig, Germany. TIBTECH JANUARY 1999 (VOL 17)

expression of heterologous genes and protect them from inactivation by the flanking chromatin4. The S/MAR effects are distinct from those of enhancers because, in the absence of chromosomal integration during transient assays, S/MARs have no influence on expression5–7. Controlled gene expression is of major importance for functional-genomics research, the production of growth-inhibitory products and complex metabolic engineering. This can be achieved by using regulatable promoters that respond to medium additives or growth conditions. The first recombinant, regulated geneexpression system developed in mammalian cells is based on the Escherichia coli lac operator–repressor interaction: the lac repressor inhibits transcription by binding to lac operator sequences that are inserted between the TATA box and the transcription start site of a mammalian promoter; transcription is activated by the inducer isopropyl-b-D-galactopyranoside, which induces the release of the lac repressor from its operator8. Because the regulatability of transgene expression using the lac system was relatively poor in most industrially relevant cell lines, most currently used regulated gene-expression systems rely on promoter–transactivator combinations (Fig. 1). For example, efficiently regulated, tetracycline-responsive gene expression has been achieved by the combination of a transactivator based on the prokaryotic tetracycline repressor and a promoter containing multiple binding sites for this transactivator9. In some cell lines, very high transcription rates have been achieved, as well as ranges of expression spanning five orders of magnitude9. In addition to the tetracyclinedependent transactivator, an inverse system that is tetracycline repressible has been developed10. In this system, tetracycline is, in practice, replaced by nontoxic analogues such as doxycyclin. The level of the transactivator plays a major role in the transcriptional activity of the tetracycline-dependent promoter, but a number of cell lines expressing tetracycline-regulatable transactivators are commercially available. In addition, a bidirectional promoter has been constructed that enables the regulated expression of two genes simultaneously11. Other regulatable transactivators that are widely used are steroid-hormone receptors. In the absence of hormones, these proteins form an inactive complex with heat-shock proteins, but ligand addition induces the transcriptional activation of the target promoters.

0167-7799/99/$ – see front matter © 1999 Elsevier Science. All rights reserved. PII: S0167-7799(98)01248-7

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A A A

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Preg Figure 1 Artificial transactivator–promoter combinations. These are the most efficient transcription systems available and allow highly effective regulation of gene expression. A generic transactivator (T) consists of a heterologous DNA-binding domain (to avoid unwanted activation of endogenous genes), a regulatory-substance (A)-binding domain and a transcriptional-activation domain. The transactivator gene is transcribed from a constitutively active promoter (Pcon) and its activity is regulated by an exogenously added substance. The target promoter (Preg) contains multiple binding sites for the transactivator upstream of an optimized minimal promoter, from which the product gene is transcribed.

Flp-driven intrachromosomal recombination Chromosomally integrated construct

Targeting vector

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Flp-driven recombination with plasmidal DNA Product

Figure 2 Heterologous recombination systems. These allow the exchange of expression cassettes at previously marked loci. A gene encoding an easily detectable marker protein flanked by recombination sites is integrated at random sites into the genome of the producer cells. To reflect the intrinsic activity of the locus, the integration must occur in single copy, which is preferentially obtained by transduction with retroviral vectors. Recombination sites can be inserted into the long terminal repeats (LTRs) of these vectors. Cells with the optimal expression pattern are identified and isolated. Expression of a heterologous recombinase (as an example, the yeast Flp recombinase is shown) catalyses the exchange of the chromosomally integrated DNA between the recombination sites (here FRT, a 13-bp inverted-repeat sequence separated by an 8-bp spacer sequence) with a product gene flanked by the same recombination sites. Improvements of this basic procedure avoid the subsequent deletion of the product gene by recombination of the flanking recombination sites with each other by using two different recombination sites. Deletion of the marker gene is an efficient but unwanted side reaction that can be selected against by including an incomplete drug-resistance marker gene in the 39 LTR. The missing gene fragment is supplied by the product cassette, such that only precise recombination events render the cell drug resistant.

Hormone-dependent transcription activators have been constructed by fusing hormone-binding domains of steroid receptors to DNA-binding domains of

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unrelated heterologous proteins12. DNA-binding domains from the yeast Saccharomyces cerevisiae Gal4 protein have been used in combination with the hormone-binding domain of the mammalian oestrogen receptor, the fusion product of which can be induced by oestradiol13. Analogously, a progesteronereceptor fusion protein has been produced that can be activated by RU 486 at concentrations much lower than those required for antiprogesterone activity14,15, as has a system based on the heterodimeric insect ecdysone receptor, which can be induced by the synthetic ecdysteroid compound ponasterone A or muristerone A (Ref. 16). The major drawback of isolating stable, inducible or constitutive high-level producer cell lines is the timeconsuming selection procedure. In order to ease the isolation of cell clones with the desired characteristics, a number of sophisticated techniques have been developed. Controlled, targeted chromosomal-integration expression Low-copy integrated genes are generally expressed more stably than the multicopy genes usually obtained with the calcium coprecipitation method17. To select transfectants with low-copy-number integrated genes that nevertheless are expressed at high levels, a neomycin-phosphotransferase gene with a mutant translation-initiation site has been used18. Alternatively, retroviral vectors can be employed to achieve singlecopy integration. In principle, if one of the rare sites on the chromosome that allow high gene expression from a single integrated gene copy has been identified, this locus can be used for high-level expression of any desired transgene. With the exception of mouse embryonic stem cells, which allow the insertion of DNA fragments by homologous recombination, the integration of transfected DNA into mammalian genomes is unpredictable and occurs almost exclusively by non-homologous (illegitimate) recombination. Therefore, for targeted transgene integration into the mammalian chromosome, a two-step approach has been developed. First, a suitable chromosomal location with high transcriptional activity must be identified by classical transfection and subsequent screening. The screening vectors used for finding these transcriptionally highly active sites contain selection markers and reporter genes flanked by artificial recombination sites such as the yeast Flp recombination target (FRT) sites, or the bacteriophage P1 loxP sites, which are recognized by the Cre recombinase. The DNA between the recombination sites can then be precisely replaced with a second cassette encoding the expression unit of the gene to be expressed19. Unwanted excision of the insert by recombination of the flanking sites can be suppressed by using two FRT sites with different nucleotide sequences20,21. Cassette exchange is made considerably easier by using a counterselection procedure (Fig. 2). An extremely effective selection procedure for correct cassette replacement uses an initiation-codon-deficient drug-resistance marker in the first selection construct. The missing in-frame initiation codon is then introduced with the second expression cassette. Only cells that have TIBTECH JANUARY 1999 (VOL 17)

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recombined precisely become drug resistant22. The tedious and time-consuming work to improve this approach is compensated by the possibility of reusing such stable, high-level producer cell lines to express any other gene of interest. Multicistronic expression Although the need for screening procedures cannot be completely removed when generating an ideal producer cell line, it can be considerably reduced by linking the expression of the product to the expression of a selectable marker. The correlation between selectable-marker activity and the amount of product secreted depends on the method used. Cotransfection of the two genes on different plasmids is the least accurate procedure but can be improved by expressing the product and the marker gene from the same plasmid. The most stable coupling is obtained when both genes are expressed from the same promoter. Two separate reading frames can be translated from a single mRNA by taking advantage of an internal ribosomal entry site (IRES) to direct translation initiation of the second cistron. Such artificial genetic configurations have become standard for coordinated, dicistronic geneexpression systems in mammalian cells, for example, to link the gene of interest transcriptionally to a selection marker or to express two-component proteins such as immunoglobulins at defined ratios23–29. Most of the IRES elements used for dicistronic expression originate from poliovirus or encephalomyocarditis virus, but other IRES sequences can be used as well30,31. The efficiency of IRES-mediated translation initiation varies greatly between different IRES elements and is also dependent on the host cell line used31. Coordinated expression of two genetic traits can also be achieved by gene fusions32 or by differential splicing33, but only IRES-mediated translation can be extended beyond dicistronic configurations (Fig. 3). A novel family of expression vectors using three and four cistrons have been constructed for coordinated, constitutive or adjustable high-level expression of three independent genes in mammalian cells34–36. Multicistronic expression has enabled the development of a number of important procedures, including: the coupled expression of multicomponent and multisubunit proteins; one-step transfection, selection and maintenance of difficult-to-express genetic configurations; the cloning of good producer-cell lines by introducing a selection marker into the last cistron (auto-selective systems)33; the design of complex regulatory circuits such as positive-feedback regulatory systems for one-step regulated gene expression in mammalian cells33; and multigene metabolic engineering of production cell lines using sense, antisense or ribozyme technology. The applicability of multigene expression cassettes in genetic immunization is currently under investigation.

methods to isolate these rare high-level producer cells include the use of two selection markers38 instead of a single selection marker39. In Chinese hamster ovary (CHO) cells, the standard procedure for achieving maximal expression is a very time-consuming procedure that can last for months and includes a gradual increase in the selection pressure for a cotransfected selection marker such as the dihydrofolate reductase or the glutamine synthetase. A more rapid method to isolate high-producing clones involves the use of plasmid vectors that encode green fluorescent protein (GFP) as a screening and fluorescenceactivated-cell-sorting marker. Alternatively, efficient expression can be enforced by employing selection markers with loss-of-function mutations. It has been repeatedly shown that the lower translation efficiency of such mutated markers is compensated for by the selection of chromosomal integration sites with overall high transcription activity28,40. Sophisticated screening procedures have been used to monitor the product level directly (Fig. 4). A relatively simple and inexpensive method relies on the local transfer of a constitutively or inducibly secreted product from a colony of producer cells through an agarose overlay onto a filter. The bound product can then be detected immunologically25,41–43. At the single-cell level, the determination of productivity is not trivial and requires the association of the individual secreting cell with its product. This has been achieved by tethering antibodies to the cell membrane of producer cells. In the low-permeability medium, the product accumulates in the proximity of each producer cell and is captured by the antibody, so that each cell becomes associated with a certain fraction of its own product. For sorting, the cell-associated product can be labelled with magnetic beads or with a second antibody44. An alternative method to keep the product associated with individual producer cells is to embed the cells in gel droplets. The product is caught by antibodies that are linked to the gel matrix. For quantification, the droplets are labelled with a second antibody. Beads containing the cells with the highest expression level can then be selected using flow cytometry45–48. Controlled-proliferation technology Cell growth and division are of fundamental importance for multicellular organisms, especially in the first developmental phase, when cell proliferation is

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Advanced selection and screening procedures Even when using multicistronic expression vectors for stable transfections, individual clones express recombinant proteins at highly variable levels37, and only a small fraction (generally ,1% of the clones initially obtained) stably express high amounts of the desired product protein. Improvements in various TIBTECH JANUARY 1999 (VOL 17)

P

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Figure 3 Multicistronic constructs express several reading frames in a defined ratio. The first cistron is translated similar to most cellular mRNAs in a cap-dependent initiation process (ci), while downstream reading frames must be preceded by an internal ribosomal-entry site (IRES) to allow cap-independent internal translation initiation (ii).

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a Filter immunoassay (i)

(ii) (iii)

b Secretion-capture report web (i)

(ii) P

c Cell-surface affinity matrix (i) P

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Figure 4 Specialized methods to isolate cells with high expression levels. In the simple and inexpensive membrane-immunoassay method (a), cell clones are covered with a thin agar overlay (i) onto which a proteinbinding membrane is placed (ii). The product diffuses locally through the agar and binds to the membrane, and is detected by labelled antibodies or another appropriate detection method (iii). The ‘secretion-capture report web’ (b) is a derivatized agarose bead into which single cells are embedded (i). The secreted product (P) is captured by biotinylated antibodies (black) that are coupled with avidin (small circle) to biotinylated agarose (ii). For detection, the agarose bead is soaked in a solution containing a labelled second antibody (grey). In the cell-surface affinity-matrix method (c), the capture antibody is coupled to the membranes of biotinylated cells and the product kept in the vicinity of the cell by a special low-permeability medium.

essential for growth of the organism. However, when terminal differentiation and the growth phase are completed, proliferation control becomes the dominant aspect of the genetic stability of higher organisms. Despite the growth-arrested state, the cells produce and secrete proteins continuously during the lifetime of the organism. In standard biotechnological production processes, transformed, permanently proliferating animal cells are used, which eventually die of nutrient or oxygen depletion. The requirement for cell growth to clone and propagate producer cell lines to high densities, and reports of a positive correlation between growth rate and product formation49–52, stimulated efforts to replace growthfactor-containing animal serum with chemical media

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containing defined proteins37,53 and genetically engineer the mammalian cell lines to permit growth in low-protein or protein-free medium54–56. In spite of successful efforts to enhance cell growth, proliferationbased cell-culture technology is hampered by the fact that growth beyond a desired cell density causes nutrient limitation, accumulation of toxic compounds and, eventually, cell death accompanied by the release of hydrolytic enzymes, which can degrade the product and decrease product quality. However, as in the natural situation, an ideal production process would include proliferation control to allow cells to grow rapidly to high cell densities, followed by a proliferation-inhibited production phase in which the cells devote their metabolic capability to the formation of product. Higher productivity of proliferation-inhibited cells was indeed observed with antibody-producing hybridoma cells57–59. However, as the proliferationinhibited state was achieved by starving the cells for an essential energy source or the addition of DNAsynthesis inhibitors such as thymidine, hydroxyurea, transforming growth factor b (TGF-b) or genotoxic agents such as adriamycin, or with temperaturesensitive mutant cells at the non-permissive temperature57,59,60, such processes or additives reduce cell viability and are therefore less suitable for long-term production systems. The first attempt to control cell proliferation by the genetic engineering of a baby hamster kidney (BHK) cell line was based on an oestrogen-regulated fusion between the interferon-responsive factor 1 (IRF-1) and the oestrogen receptor61–63. IRF-1 is a DNA-binding transcription activator that accumulates in cells in response to interferons and has antiviral and antiproliferative activities25,59,61. The induction of recombinant protein production can be achieved in IRF-1-arrested cells by placing the transcription of the corresponding gene under the control of an IRF-1responsive promoter. In addition, the expression of oestrogen-responsive IRF-1 in a dicistronic configuration with the selection marker stabilizes the growth control in a way that allows several cycles of growth and growth-arrested states to be performed with the same culture63. In another effort, the cyclin-dependent-kinase inhibitors p21 and p27, and the tumour-suppressor gene p53 were used to arrest CHO cells reversibly at the G1 phase of the cell cycle65. p53 functions as a key regulator at the interconnection of the cell-cycle and apoptosis regulatory networks, and either affects apoptosis or induces p21, to allow repair and survival66. Overexpressing any one of the three genes leads to a repression of cyclin-E–CDK2-complex phosphorylating activity and prevents entry into S phase66. The G1 phase of the cell cycle is expected to be the most suitable for controlled-proliferation technology as it is the phase in which a cell checks its physiological state and can arrest to replenish its metabolic supplies and repair genetic defects66. CHO cells stably transfected with multicistronic p21-, p27- or p53-containing tetracycline-repressible expression vectors show different responses. The regulated overexpression of p27 leads to growth arrest with no signs of apoptosis and up to 15-times-higher specific TIBTECH JANUARY 1999 (VOL 17)

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productivity67. The continuous overexpression of p53 [even in its mutated, apoptosis-deficient, form (p53175P)] leads to rapid cell death68. No clones could be isolated that showed proliferation arrest after induction of p21 expression. Making use of tetracyclineregulated tricistronic expression68, the dicistronic SEAP–p21 expression unit has been extended with a cistron encoding the differentiation factor c/ebp a (CAAT-enhancer-binding protein a) that both induces and stabilizes p21 (Ref. 68). With this construct, sustained growth arrest for several weeks and a 10–15-fold higher specific productivity has been achieved compared with control cells67. A second example for controlled-proliferation technology is based on a p27–bcl-xL-encoding tricistronic expression unit. CHO cells stably transfected with this construct showed a 30-times-higher expression level than did the controls. Multigene metabolic engineering of the cell cycle has thus proved to be effective in achieving a difficult-to-attain cell-culture state and enhancements in the expression of a heterologous secreted glycoprotein. Transient expression As an alternative to the cumbersome and timeconsuming selection procedure of stable cell lines, transient transfection strategies can be used for the rapid and efficient production of small quantities of recombinant product protein. Cell lines that express a particular transcriptional activator can drive high-level expression from specific promoters that contain binding sites for the respective transactivator. For example, replicating expression vectors have been developed for the human 293 cell line that constitutively expresses the adenoviral E1A transactivator to enhance transcription, concomitant with the EBNA antigen and Epstein–Barr virus replicon for episomal plasmid maintenance69. In addition, COS monkey cell lines are still used for extrachromosomal plasmid replication and efficient gene expression from SV40 origin promoters70–72. Many of the currently available viral vectors can be used for transient gene expression in mammalian cells. Vectors derived from Papova viruses, Epstein–Barr virus, adenovirus or alphavirus replicate episomally and often reach very high copy numbers, resulting in very high, transient expression levels. However, their productive time span is limited owing to the loss of the expression construct or the death of the host cell. Alphavirus-derived vectors have been developed for protein production73, as they can infect non-dividing cells and have a broad host range. The alphavirus genome consists of a 12 kb positive-strand RNA molecule, which is translated to produce a replicase immediately after infection. Replication signals at both ends of the RNA are required for the amplification of the viral genome that takes place in the cytoplasm until the infected cell dies. Negative-strand RNA is produced that serves as a template for the synthesis of both full-length genomic RNA and up to 105 RNA molecules of a subgenomic transcript encoding structural proteins. These structural genes can be replaced by heterologous genes, and the recombinant viral RNA then packaged with the help of a coexpressed wild-type virus or a packaging cell line expressing complementing structural proteins. TIBTECH JANUARY 1999 (VOL 17)

Current alphavirus-based expression vectors allow nonviral gene transfer using either in vitro-transcribed RNA or, alternatively, DNA vectors from which the viral RNA is transcribed under the control of a heterologous promoter. The potential risk of a selfreplicating infectious virus is reduced by a conditionally activated Semliki–Forest-virus expression system in which a cleavage-deficient spike protein is activated by the exogenously added protease chymotrypsin74. Vectors based on vaccinia virus have emerged as the most efficient transient expression system15. However, at present, only a few laboratories have the expertise required to handle this system. Among other difficulties, the virus might induce changes in the posttranslational system of the host cells, which could lead to premature cell death. Therefore, vaccinia-virus and alphavirus vectors have been used mainly in recombinant protein production for research purposes and in animal vaccination75. Although all of the above-mentioned viral vectors combine the two prerequisites for strong transient expression – efficient gene transfer and high episomal copy number – these properties have also been recently achieved by refinements in classical transfection technology. Improvements in DNA preparation and transfection protocols now enables the production of grams of recombinant proteins in transient-transfection batch-fermentation processes76,77. Apoptosis engineering Apoptosis is a genetically determined program of active cell death. Despite its fundamental importance for multicellular life, apoptosis is an undesirable phenomenon in biotechnological production processes, which are often limited by rapid cell death in the decline phase of a culture. Many commercially important production cell lines are sensitive to apoptosis, including hybridoma and myeloma cell lines78,79; others, such as HeLa or HL-60 cells, block cellular proliferation in response to nutrient limitation or genotoxic stress rather than initiating apoptosis, and allow cells to replenish their metabolic precursors or repair DNA damage80. There are three fundamental approaches to suppressing apoptosis in cell-culture processes81,82: (1) elimination of nutrient deprivation by feeding strategies; (2) use of chemical medium additives to block apoptosis pathways; and (3) metabolic engineering using antiapoptotic survival genes. Suboptimal concentrations of amino acids, particularly essential amino acids, can activate apoptotic signalling pathways78,83. The addition of a single amino acid has recently been shown to rescue hybridoma cultures from starvation-induced apoptosis84. Medium additives that prevent apoptosis include chemicals that inhibit the activity of the endonuclease responsible for DNA cleavage81, antioxidants such as vitamin E (which can prevent apoptosis induced by free radicals85) and pseudosubstrates for caspases, which block the activity of these apoptosis-inducing proteases86. Apoptosis is induced by a wide variety of stresses typically encountered during normal bioreactor operation87. These include nutrient and serum limitations, accumulation of toxic compounds and metabolites, oxygen deprivation, and hydrodynamic stresses.

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Mammalian cells can be protected against stressinduced apoptosis by genetic engineering to overexpress survival genes such as Bcl-2 (Refs 60, 80). Enhanced Bcl-2-mediated survival has also been reported for Sindbis-virus-based expression systems89. The level of protection varies between different cell types and cell lines and, in most cases, Bcl-2 overexpression cannot prevent cell death, but it can extend cellular lifetimes and lead to increased production59,88,90–93. Initial attempts to complement the action of Bcl-2 using concomitant overexpression of the antiapoptotic genes bag-1, bcl-xL or the adenoviral E1B-19K gene were encouraging and showed that the protective effects of individual genes were equal or even additive when, for example, Bag-1 and Bcl-2 were coexpressed94,95. Apoptosis engineering might become an essential tool for emerging technologies such as gene therapy and tissue engineering. Glycosylation: a target for post-translational metabolic engineering Although gene dosage, transcription levels and translation rate are key parameters for optimizing heterologous protein production in mammalian cells, posttranslational processes have only recently been perceived as metabolic bottlenecks and potential targets for improving the performance of mammalian cell culture. Post-translational modification of a product protein includes various potentially rate-limiting interactions with numerous chaperones and enzymes in the cytoplasm, the endoplasmatic reticulum and the compartments of the Golgi apparatus. Studies in mammalian cells96–98, yeast99,100 and insect cells101,102 have improved the protein productivity by augmenting the post-translational capabilities of these cell systems. One of the main post-translational events is the attachment of complex sugar structures to the majority of secreted proteins. Glycoproteins mediate many diverse functions65,103–106 and the glycosylation pattern of secreted proteins influences their activity as well as their clearance from the body. Certain amino acid residues serve as specific glycosylation sites as the polypeptide chain moves from the endoplasmic reticulum through the Golgi apparatus. Two major types of glycosylation are commonly found in eukaryotic cells: N- and O-linked glycosylation, with carbohydrates attached to the amide group of Asn–X–Ser/Thr sequences or the hydroxyl groups of Ser or Thr residues, respectively. The final glycosylation pattern of a protein is dependent on many parameters including: (1) the polypeptide chain; (2) the host cell and its chromosomal set of glycosyltransferases and glycosidases; and (3) the environment of the host cell. Different polypeptides expressed in the same host under similar conditions can be glycosylated very differently, showing that the polypeptide itself exerts a controlling influence on its own glycosylation107. A technologically relevant example shows that a single amino acid substitution in tissueplasminogen activator (tPA) can create a new glycosylation site and even change the glycosylation pattern of a different native site. Although this tPA glycovariant showed a ten-times lower plasma clearance than wild-type tPA, its physiological activity decreased to one third of the wild-type level, a phenomenon that

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could be alleviated by removing the native glycosylation site108. Even with a given protein product and a welldetermined host system, the oligosaccharides present on secreted glycoproteins are built up in a complex cascade of sequential enzyme-catalysed reactions and intracompartmental transport processes, often with multiple enzymes acting on common substrates to give alternative oligosaccharide products109. A particular glycoprotein produced in a given host system is a population of different glycoforms. These differ: (1) in the percentage by which a particular amino acid residue is glycosylated; (2) in high-mannose or complex glycoforms in N-linked oligosaccharides; (3) the number of terminal residues (antennae); (4) in bisecting N-acetylglucosamine (GlcNAc); (5) in saccharide–saccharide linkages; and (6) in the percentage of oligosaccharide ends that are sialylated. Glycoforms can significantly differ in their activity107,110, physicochemical properties106,110,111 (solubility, stability, folding and secretion), pharmacokinetics (clearance of a protein pharmaceutical from the blood stream104,111–114, targeting, immunogenicity, antigenicity and stability104. Data on the functional implications of attached oligosaccharides are only available for a very limited number of glycoproteins. A few general principles of carbohydrate structure–function relationships are known. A central concern of biopharmaceutical manufacture is the absence of sialic (neuraminic) acid on the termini of complex carbohydrate structures, which results in more rapid clearance, as does the presence of high-mannose oligosaccharides. In addition to the protein itself, glycosylation is dependent on the host cell and the culture conditions115. Commonly used bacterial hosts do not glycosylate their proteins; yeasts glycosylate with highmannose oligosaccharide structures that are not suitable for injection into humans. Similarly, baculovirus and plant expression systems synthesize carbohydrates that are undesirable for pharmaceutical applications. It is the capability of mammalian cells to accomplish glycosylation in a manner compatible with human applications that has created a special niche for them in the biotechnology industry. However, even mammalian cells are far from being perfect in their glycosylation. Current research focuses on strategies to modify targets and oligosaccharides on particular glycoprotein pharmaceuticals to produce glycoforms with enhanced therapeutic potential. Methods developed to manipulate glycosylation include drugs to inhibit glycosylation as well as inhibitors of glycosylation processing116. Many of these reagents are toxic and sometimes alter glycosylation patterns rather inefficiently. Other strategies focus on random mutagenesis for the generation of host cells with altered glycosylation characteristics. Screening is facilitated by a lectin-based (lectins are toxic carbohydrate-binding proteins) counterselection of wild-type cells and an enrichment for host cells with altered glycosylation patterns of the surface proteins103. However, such a selection of loss-of-function mutations in the glycosylation pathways often results in incomplete or truncated carbohydrate structures. The use of recombinant DNA technology for the metabolic engineering of glycosylation in mammalian TIBTECH JANUARY 1999 (VOL 17)

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cells aims to extend the hosts’ oligosaccharidebiosynthesis capabilities by introducing genes encoding heterologous carbohydrate-synthesis enzymes. At present, most glycosylation-engineering strategies have considered enzymes involved in or near the terminal steps of complex oligosaccharide biosynthesis117–120. Two main metabolic-engineering strategies are being followed; glycosylation activities can be increased based either on: (1) gene activation; or (2) introducing transcriptionally controlled glycosylation genes. Conversely, antisense and other methods can be used to block undesired glycosylation. Only a few glycosylation enzymes have been cloned and are available for metabolic engineering120. CHO cells lack a functional copy of the a2,6-sialyltransferase gene (2,6 ST)121. This defect of CHO cells, relative to human cells, can be complemented by the expression of a cloned 2,6 ST, which results in the production of both a2,3- and a2,6-linked sialic acids on coexpressed tPA (Ref. 117). The glycosylation pattern of heterologous pharmaceutical proteins was changed in BHK cells when the normally absent a2,6-sialyltransferase and a1,3-fucosyltransferase-III were overexpressed116. The metabolic engineering of glycosylation has significant potential as a means for the production of glycoproteins and to provide new forms of glycoproteins with improved therapeutic characteristics. This potential is mainly limited by the availability of cloned genes involved in the glycosylation pathways. As more components involved in glycosylation are discovered, this field will not only have an impact on further development of animal cell technology but will also allow further insights into hidden secrets of genome–cell-function relationships. Conclusion Many promising developments using genetic tools for the improvement of mammalian cell culture technology could not be reviewed here and many of the briefly mentioned techniques are still under investigation. Some of these recent developments might not be of any further use, others might become routine methods in a not-so-distant future. Genetic optimization of glycoprotein production in mammalian cells has evolved as a major multidisciplinary field of modern biotechnology in which academia, industry and medical institutions cooperate closely in order to improve current therapeutic strategies such as cell and gene therapy, and to ensure a high quality of human life. Acknowledgments Research in the authors’ laboratories on genetic engineering to control cell proliferation and to reduce apoptosis is supported by the Swiss Bundesamt für Bildung und Wissenschaft (BBW) and by the Framework IV Biotechnology programme of the European Commission. References 1 Hauser, H. (1997) in Mammalian Cell Biotechnology in Protein Production (Hauser, H. and Wagner, R., eds), pp. 3–32, W. DeGruyter 2 Kaufman, R. (1990) Methods Enzymol. 185, 487–512 3 Boshart, M., Weber, F., Jahn, G., Dorsch-Häsler, K., Fleckenstein, B. and Schaffner, W. (1985) Cell 41, 521–530 TIBTECH JANUARY 1999 (VOL 17)

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