In the last two decades, recombinant protein yields from mammalian cells in batch and extended-batch bioprocesses have increased dramatically. In general, this has been a result of the utilization of cell lines with high specific productivities, the formulation of media to allow the suspension cultivation of cells to high densities, a better understanding of bioprocess conditions, and the enhancement of cell viability in these high-density suspension cultures. This article discusses the contributions of these factors to yield improvement. It also describes a theoretical model to show how volumetric productivities may increase in the future. The major consequence of higher volumetric productivities is expected to be the decline in the time or volume of production runs. This outcome is expected to lead to a substantial improvement in the efficiency of protein manufacturing. 

About half of all recombinant therapeutic proteins currently on the market are produced in mammalian cells, with about 70% of these being generated in suspension culture-adapted Chinese hamster ovary (CHO) cells.

  CHO was chosen as the host for the production of tissue plasminogen activator (tPA), the first recombinant therapeutic protein from mammalian cells to gain regulatory approval. Other biopharmaceutical manufacturers continued using CHO cells because these were then considered an acceptable host system for products intended for intravenous administration in humans, but with time other mammalian cell lines such as mouse myelomas (NS0 and SP2/0), baby hamster kidney (BHK-21) cells, and human embryonic kidney (HEK-293) cells became acceptable as hosts for recombinant protein production.

  Besides the approximately 140 recombinant therapeutic proteins already on the market, there are hundreds of protein products currently in clinical trials with the majority of these being recombinant monoclonal antibodies (MAbs) produced in CHO cells. Therefore, this cell line will continue to be the workhorse of the biopharmaceutical industry for the coming decades. 

Although the abundance of candidate therapeutic proteins is good news for the biopharmaceutical industry and for public health, this success has brought its own set of problems. Undoubtedly, a major one is the so-called "capacity crunch," the expected limitation in the number of manufacturing facilities to produce the quantities of MAbs and other recombinant proteins necessary to meet the market demand in the coming years. One obvious solution is building additional manufacturing plants, but this path is both costly and slow. Alternatively, manufacturers may seek novel solutions to the most widely used type of manufacturing process—the production of recombinant proteins from suspension-adapted cell lines cultivated in large stainless-steel stirred-tank bioreactors. One avenue is developing disposable cultivation systems for part or all of the manufacturing process. Another is refining methods and technologies to allow protein production from cultivated mammalian cells to be more efficient by increasing the volumetric yields from these processes. Although this has always been one of the major driving forces for the manufacturers of biopharmaceuticals, there is now a heightened urgency for more rapid progress on yield improvement, given the sheer number of recombinant proteins in clinical trials. 

As discussed earlier, volumetric yields from recombinant mammalian cell lines have increased dramatically over the span of 20 years since the approval of tPA.

  In 1986, the industry standard for production from stable CHO-derived cell lines was a specific productivity of 10 pg protein/cell/day with volumetric yields of 50 mg/liter for batch processes that lasted up to seven days.

  By 2004, the highest reported specific and volumetric productivities were 90 pg/cell/day and 5 g/L, respectively, for an extended-batch culture lasting up to three weeks. How have these increases been achieved?  Unfortunately, the answer remains obscure because manufacturers are reluctant to reveal their insights into the generation and cultivation of recombinant cell lines. Nevertheless, it is clear that four main factors have contributed to the overall improvement in protein yields: (i) the generation of recombinant cell lines with high specific productivities, (ii) the formulation of media to support high-density cell cultivation, (iii) the understanding of bioprocess conditions for cell cultivation, and (iv) the sustained viability of cell lines in high-density batch and extended-batch cultures. We will summarize how each of these factors has supported the increase in volumetric recombinant protein productivity observed in  recent years. It is likely that manufacturers have taken different routes to establish production principles resulting in multigram per-liter-yields. 

Generating Cell lines with High Specific Productivity

Obtaining high-producing recombinant cell lines remains among the top priorities for protein manufacturers. Cell lines are generated following delivery of the gene of interest and the selection gene—on a single plasmid or on separate plasmids—into host cells by transfection. The most widely used selection markers are dihydrofolate reductase (DHFR), an enzyme that produces a cofactor for thymidylate synthetase, and glutamine synthetase (GS).

 These two markers are mainly used for selection in CHO and NS0 cells, respectively. In both cases, selection occurs in the absence of the appropriate metabolite(s): glycine, hypoxanthine, and thymidine for DHFR and glutamine for GS. Cells surviving selection are characterized by the integration of one or more copies of the transfected plasmid(s) at a single site in the cell's genome. The DHFR and GS selection markers have the advantage of supporting amplification of the copy number of the integrated DNA by exposure of the selected cells to increasing amounts of methotrexate (MTX) or methionine sulphoximine (MSX), respectively.

  Usually, no effort is made to integrate the transfected DNA to a specific site of the host cell's genome. Instead, this process is allowed to proceed randomly, and the selected cell lines are then screened for protein productivity and growth rate. The superior cell lines are also eventually analyzed for the stability of protein production over time. This strategy for cell line generation depends on the screening of a large number (100s to 1,000s) of recombinant cell lines for the desired characteristics. Strategies to increase the percentage of high-producing cell lines in the population of transfected cells include increasing the stringency of selection or amplification.

The structure of the chromatin at the site of integration is also a critical regulatory factor with regard to expression of the integrated plasmid DNA. Integration of the plasmid DNA into actively transcribed regions of the genome (euchromatin) favors high and stable expression of the recombinant gene(s), whereas integration into condensed, and therefore, transcriptionally inactive regions of the genome (heterochromatin) results in the repression of gene expression. Including cis-regulatory elements such as insulators, scaffold and matrix attachment regions (S/MARs), ubiquitous chromatin opening elements (UCOEs), and antirepressor elements near the promoter or enhancer of the recombinant gene has proven to be an effective strategy for limiting heterochromatin formation at the site of plasmid DNA integration.

  Methods to target integration of transfected plasmid DNA to transcriptionally active sites of the genome have been developed and may have been used to generate high-producing cell lines by some manufacturers.

  However, convincing evidence showing that targeted integration consistently yields cell lines with higher specific productivities and with increased long-term expression stability than those generated by random integration has not yet been published. Indeed, the recent exploitation of chromatin regulatory elements, as described above, appears to have reduced the need for targeted integration. 

The recovery of cell lines with high specific productivity also has been improved through the development of high-throughput screening tools.

  By increasing the number of candidate cell lines screened, the probability of finding a few highly productive ones is increased. The new methods are mainly based on the recovery of individual cells by fluorescence-activated cell sorting (FACS). As an example, genes for the protein of interest and a reporter protein such as green fluorescent protein (GFP) can be co-transfected into cells. If the two genes are expressed from a bicistronic mRNA having a internal ribosome entry site (IRES) for translation of the reporter gene at the 3' end of the mRNA, then cells selected by FACS for a high level of GFP are expected to be high-producers of the recombinant protein of interest. One drawback of this approach is that the intracellular accumulation of GFP may be limited by several factors and may have negative effects on cell growth and cell survival. To avoid these secondary problems, it is preferable to use high-throughput methods that directly measure the level of the recombinant protein of interest without using a reporter protein.  

Bioprocesses for High-Density Cell Cultivation in Suspension

Optimized recombinant cell lines derived from CHO and NS0 are now grown in single-cell suspension cultures to densities of 10 x 10

  cells/mL or higher during batch and extended-batch processes, a five-fold increase in cell density compared to the production processes of the 1980s. Some of these parental CHO and NS0 cells may have been genetically engineered to facilitate growth to high cell densities, but this is surely not a requirement, because such densities can be obtained in naïve cell lines, as has been observed in the authors' laboratory and by other groups.

  As an alternative to genetic host cell engineering, the parental cell line or its recombinant derivatives typically are selected from many different phenotypes of cells obtained when identifying cell lines that show characteristics applicable in bioreactor scale-up scenarios and growth under high-density situations during the production phase.  

High-density cell cultivation in suspension has also been made possible through improvements in media formulation. The media used for today's bioprocesses frequently are chemically defined and lack animal-derived components. Most protein manufacturers use in-house formulations rather than commercially available ones. Unfortunately, medium development has to be done on an individual basis for each recombinant cell line. This is a time-consuming task as the effects of multiple components must be tested alone and in various combinations. For this reason, multifactorial design has been implemented to streamline the optimization process. In addition, different formulations are necessary for different phases of a manufacturing process, each one designed for a specific phase of the process. Typically, batch and extended-batch processes can be divided into two phases, one for rapid growth to a high cell density and the other for protein production under conditions in which the cells are maintained with little or no growth. Therefore, the dilution and feeding strategies for culture scale-up and maintenance must be carefully planned, based on an understanding of the growth and metabolism of each individual recombinant cell line. The improvements in media formulation and in bioprocess control undoubtedly have contributed extensively to the prolongation of the viability of high-density cultures, but how different manufacturers have optimized their large-scale production processes can only be guessed. Furthermore, the maintenance of cells at a high density without cell growth can be achieved by different means including temperature reduction or addition of chemicals that interfere with cell physiology resulting in viability enhancing and stabilizing consequences. 

Protein production also has been increased through improvements in the robustness of recombinant cell lines so that they retain viability for long cultivation times. Increasing the resistance of cells to apoptosis probably plays a role in the prolonged production phases now observed for extended-batch cultures (i.e., up to three weeks). How increased resistance to apoptosis is achieved may differ from one manufacturer to another. There have been many published accounts of enhanced cell survival and productivity as a consequence of the overexpression of exogenously provided anti-apoptotic genes such as Bcl-2 or Bcl-xL.

  However, the gains in cell viability and recombinant protein production reported in these studies only have been modest. Therefore, it is not clear that this strategy has played a significant role in the increased viability of recombinant cell lines used in manufacturing processes. Alternatively, parental cell lines or recombinant derivatives with elevated resistance to apoptosis may have been selected, based on survival under cultivation conditions with increased cell stress. This approach may be more likely to succeed than the host-engineering strategy, given the complexity of apoptotic pathways. Moreover, host engineering by overexpression of exogenous genes may have unintended consequences with regard to other cellular functions, resulting in cells with undesirable phenotypes. The most important impact, in the opinion of the authors, is a well-balanced match of the cells' needs at specific phases of the process and optimized media and feeds provided throughout an extended-batch production phase.  

Future Improvements in Protein Yield: A Modeling Approach

To anticipate how much recombinant protein production yields may improve in the near-and long-term future, we have modeled volumetric productivities for hypothetical CHO-based production processes, using a recombinant monoclonal antibody as the product. We began by assuming a doubling time of 18 hours and a maximal cell density of 10 x 10

  cells/mL in an extended-batch process lasting up to 21 days (Figure 1). The  volumetric productivity was then determined for this process based on four different specific productivities ranging from 20 to 200 pg/cell/day. We are aware that a specific productivity of 200 pg/cell/day may be entirely unrealistic, even from a purely fundamental biological perspective. With the lowest specific productivity assumed, the volumetric productivity was about 5 g/L for a three-week production run (Figure 2).  We note that a specific productivity of 20 pg/cell/day is already a quite reasonable value, which may be the maximum achievable for certain molecules, even antibodies. By increasing the specific productivity to 100 pg/cell/day, the highest productivity yet reported for an extended-batch process, the volumetric productivity was about 10 g/L for a two-week process and about 15 g/L for a three-week process. With another doubling of the specific productivity to a theoretical value of 200 pg/cell/day, a final product titer of about 40 g/L was observed after 21 days of culture.  

Next, we increased the cell density to 20 x 10

  cells/mL for a production process lasting up to three weeks (Figure 1). Again, the specific productivities were varied from 20–200 pg/cell/day, as described above. Given these parameters, a cell line with the lowest specific productivity yielded about 10 g/L of recombinant antibody during a three-week process (Figure 3). When the specific productivity was increased to 50 pg/cell/day, final product concentrations of 10 and 20 g/L were reached with a two-week and a three-week process, respectively. By doubling the specific productivity to 100 pg/cell/day, the volumetric productivity was about 30 g/L for a two-week process and 40 g/L for a three-week process. Yields of 80 g/L were obtained for a 21-day process when assuming a specific productivity of 200 pg/cell/day. At this specific productivity, a titer of 20 g/L was achieved in a theoretical process lasting seven days (Figure 3). 

Based on the modeling results shown here, it is clear that doubling the volumetric productivity from the current record level reported (5 g/L) to 10 g/L will not be difficult to achieve for cultures at a density of 10 x 10

  cells/mL and with cell lines with specific productivities in the range  of 50–100 pg/cell/day. By doubling the cell density to 20 x 10

  cells/mL, the 10 g/L final product yield can be reached in about two weeks for a cell line with a specific productivity of 50 pg/cell/day, in about five days for one with a specific productivity of 100 pg/cell/day, and in about three days at a specific productivity of 200 pg/cell/day (Figure 3).  

In view of the widely known capacity limitations in downstream processing, mostly with respect to the first product capture step, it is questionable whether higher product concentrations (than 2–5 g/L) in harvest fluids are even desirable.

  This may change in the future if radically new product recovery principles become available and acceptable. However, as the above model calculations show, shortening of production runs will be a very useful approach to economize the valuable cell-culture infrastructure without reducing overall product yield derived from a manufacturing facility. More individual runs for a given product in a given timeframe can liberate the facility for other products to be subsequently made in the same facility. Of course, if capital investment is required for construction of a new facility by small and mid-sized biotech companies, the obtainable yields from cell-culture processes will provide options for the construction of smaller facilities, thus reducing the financial burden. As an example, a cell line at a density of 10 x 10

  cells/mL and a specific productivity of 50 pg/cell/day will yield 10 g/L in a three-week process. If the cell density is doubled, then the same amount of recombinant protein can be produced in half the volume in three weeks.  

In the model presented here, we have assumed that protein quality, particularly the extent of glycosylation and aggregation, remains the same no matter what the specific productivity or the cell density is. However, as experience has shown, such an assumption is surely an oversimplification. Typically, careful adjustments to media compositions and numerous process details must be implemented to maintain product equivalence when extending or shortening run times and or changing the scale of operation in the final production vessel.   

Over the last 20 years, the yields for bioprocesses performed in single-cell suspension cultures of mammalian cells have increased dramatically. It remains to be seen whether the physical properties of secreted proteins, in terms of solubility or in interactions with other cell-derived components, may eventually limit the  rise in product concentrations in cell culture media. Undoubtedly, the overall cell densities in reactors will rise further and provide the most promising approach for further improvements in final product concentrations. Even at the highest reported cell densities of 10–15 x 10

  cells/ mL,  the total biomass represents less than 5% of the culture volume. The other 95% of the volume is composed of the liquid medium and the soluble components therein. Higher cell densities will depend on better basic and feed media. This vast area of research has not obtained the visibility and recognition of novel vector systems and productivity-enhancing DNA elements. Whether specific productivities of 50–100 pg/cell/day can, in the future, be reached routinely for the majority of recombinant proteins is to be found out, but surely this goal depends on careful screening and better and more efficient tools to identify such superproducers that also have the multitude of characteristics necessary to survive under scale-up and production conditions. 

 is a professor, all at the Laboratory of Cellular Biotechnology, Lausanne, Switzerland. Wurm is also the founder and chief scientific officer of ExcellGene SA, Lausanne, Switzerland, +41.21.693.6141, 

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 4. Kwaks TH, Otte AP. Employing epigenetics to augment the expression of therapeutic proteins in mammalian cells. Trends Biotechnol. 2006 Mar;24(3):137–42. 

 5. Baer A, Bode J. Coping with kinetic and thermodynamic barriers: RMCE, an efficient strategy for the targeted integration of transgenes. Curr Opin Biotechnol. 2001 Oct;12(5):473–80. 

 6. Browne SM, Al-Rubeai M. Selection methods for high-producing mammalian cell lines. Trends Biotechnol. 2007 Sep;25(9):425–32. 

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Articles

A discussion of past achievements and future expectations of recombinant protein production yields from mammalian cells.

Recombinant Protein Production Yields from Mammalian Cells: Past, Present, and Future

Culture media development continues to be an area that offers the potential to dramatically improve the productivity of biopharmaceutical manufacturing processes. Having a well-defined media formulation optimized for maximum protein production can significantly improve product titers, thereby reducing costs and improving efficiency. In this article, we discuss the historical aspects of media development from the initial dominance of serum as a supplement in cell-culture media to the current trend toward serum-free, chemically defined, protein-free media customized for particular cell lines. We will also examine the advantages of developing a robust serum-free cell-culture medium containing defined animal-free protein supplements.

The past three decades have seen major developments in cell culture technologies in response to the increasing demand for approved biopharmaceuticals. Various approaches have been used to generate cell culture processes with the desirable traits of high titer, robustness, and improved scale-up efficiency. Key focus areas for upstream process development and optimization include choice of cell line and clone, expression vector design, medium optimization, and bioreactor conditions. This article examines approaches to media development in the biopharmaceutical industry over time, the impetus for change, and directions for the future. 

The first application of cell culture technology was for vaccine production in the 1950s. Traditionally, embryonated chicken eggs were used for vaccine production. Because of increased demand for vaccines, there was a need for new technologies that could produce large quantities. The first mammalian cells used to generate a commercial product were monkey kidney cells, which were used to produce the polio vaccine.

 These were an attached cell line and required serum for growth. Further advances occurred during this time with the commercial production of an inactivated foot-and-mouth disease vaccine in baby hamster kidney (BHK) cells. This cell line was cultured in Eagle's medium, supplemented with 5% adult bovine serum up to a scale of 5,000 L.

  Eagle's minimal essential medium, the first basal synthetic medium, contained a mixture of amino acids, vitamins, cofactors, carbohydrates, and salts to support cell growth. However, supplementation was also required, generally by addition of animal serum. Another major advancement in the use of mammalian cell culture for the production of biologics for human use came with the US Food and Drug Administration's acceptance of the use of continuous cell lines. Continuous cells lines such as Chinese hamster ovary (CHO) and myeloma cells (NSO and SP2/0) meant cells could be cultured in suspension and were readily scalable, allowing for increased product yields. 

Bovine serum has been the supplement of choice during the development of general cell culture. Serum has been shown to provide all the essential nutrients for cell growth and productivity, including macromolecular proteins, low-molecular-weight nutrients, anti-oxidants, and carrier proteins for water-insoluble components, and is known to contain anti-apoptotic factors.

  Serum also contains a high concentration of albumin, which can protect cells from stress factors generated under bioreactor conditions such as pH nutrient fluctuations and shear forces, as well as a number of transport functions.

  Although serum delivered many benefits for the industry, serum also has important drawbacks, such as batch-to-batch variability, leading to a lack of process and product consistency, complex downstream processing to remove serum contaminants from the final product, and fluctuating costs. Even more important, the transmissible spongiform encephalopathies (TSEs) and other adventitious agents in bovine serum resulted in a strong regulatory drive for the biopharmaceutical industry to eliminate serum and other animal-derived components from the manufacturing process.

  A 1999 report by Wessman and Levings indicated that as much as 20–50% of commercial fetal bovine serum was virus-positive.

  These regulatory constraints led to the development of defined media supplements and the industry's move away from the use of serum to a serum-free medium. 

Initially, serum-free medium was introduced into biopharmaceutical manufacturing because of the cost and process constraints of serum. Serum-free medium (SFM) was first used in mammalian cell culture by Ham in 1965. Much of this early work involved anchorage-dependent cell lines.

  To achieve growth-promoting effects similar to those of serum, SFM often was supplemented with animal-derived components that replaced the role of serum, such as transferrin, albumin, insulin, and other biological extracts. Serum supplementation continues to be used in many early stage research applications, clonal development, and vaccine production.  

Establishing more defined media regimes reduced the problems associated with the batch-to-batch variability seen with serum and allowed more consistent product and process control with simpler purification and downstream processing strategies. However, having more defined media often resulted in extended cell adaption times, reduced growth rates, and decreased product titers, all of which increased the costs of manufacture.  

The most common serum-free supplements incorporated into a basal medium have been growth factor sources, transferrin, hydrolysates, and albumin. Even today, some of these supplements are animal-derived (transferrin, albumin), contain animal components, or are ill defined and are still used in manufacturing processes because of the lack of safe alternatives. The regulatory issues of possible contamination of the final product with adventitious agents and ill-defined processes therefore remain. Chemically defined SFM are commercially available, but for some cell lines it has not been possible to design a robust, animal-free, and chemically defined medium that will perform as well as serum. Often, different cell lines and clones exhibit a large degree of variability in their nutritional requirements to achieve optimal growth and performance, resulting in a lengthy and costly media development program.  

The remainder of this article will discuss four media supplements commonly used in the biopharmaceutical industry and recent advances in their development. 

Regulatory agencies have encouraged the development of serum-free media devoid of any animal-derived components to avoid the risk of introducing adventitious agents such as viruses and prions. Protein hydrolysates have been shown to have beneficial effects on cell growth and productivity and are a relatively effective alternative to the use of serum. Peptones derived from bovine milk or animal tissues, such as Primatone RL (MP Biomedicals, Irvine, CA), are capable of supporting a number of different cell lines in the absence of serum.

  However, non-animal hydroly-sates from microorganisms such as yeast

  are being investigated as supplements for supplying the nutritional requirements of mammalian cells in culture. 

Although plant-based hydrolysates have been shown to promote growth and productivity,

  the industry has not fully embraced hydrolysates as a serum substitute. Primarily this has been because of a lack of chemical definition and lot-to-lot variability leading to process and product inconsistencies. Currently, manufacturers of hydrolysate supplements are addressing these concerns through novel enzymatic techniques and more refined processing to produce a more consistent product.

Albumin, in the form of bovine or serum albumin (BSA, HSA) is commonly used in cell culture media formulations for nutrient transport. SFM is frequently supplemented with serum albumin as a carrier for fatty acids, lipids, amino acids, and trace elements. Additional advantages of albumin as a cell culture supplement include its ability to bind toxic components present in culture and protect against mechanical damage such as shear stress in agitated cell culture systems. The successful replacement of BSA or HSA in SFM with recombinant forms of albumin or synthetic compounds such as pluronic has been achieved, however, requirements for albumin vary depending on the cell line. For example, the myeloma cell line, NSO, lacks the functional pathway for cholesterol synthesis and therefore requires cholesterol. Albumin has been used as a carrier of cholesterol, although cyclodextrins have been used as alternative carriers of cholesterol and other lipids in culture media. 

In recent years, a variety of recombinant animal-free forms of albumin (rHA) have become commercially available. Up and coming areas requiring commercial cell culture processes, such as advanced tissue and stem cell therapies and regenerative medicine, are areas where rHA is likely to prove a compliant and consistent alternative to current albumin sources.

  Recombinant albumin, such as CellPrime rAlbumin AF-S (Millipore, Billerica, MA), and Recombumin (Novozymes, Bagsvaerd, Denmark), produced in 

 has been shown to be structurally identical to native albumin. Safety, tolerability, pharmacokinetics, and pharmacodynamics also have been studied, showing equivalence to native human albumin.

Although cells may be adapted to grow in media devoid of growth factors, growth factor supplements still are essential for the growth of many cells in culture. Insulin traditionally has been used as a mitogen and also is involved in glucose amino acid uptake, lipid metabolism, and DNA synthesis.

  Recombinant insulin has been available since 1982 (Genentech, Eli Lilly) and is the most universal supplement in SFM. Although insulin is the growth factor of choice, it is required at supra-physiological concentrations (2–10 mg/mL) to support cell growth and viability under culture conditions.

  It is widely accepted that insulin action is primarily through the activation of the IGF-I receptor (IGF-IR) rather than its own insulin receptor (IR).

An insulin-like growth factor analog, LONG R

 IGF-I (Novozymes) has been developed that acts directly at a much higher potency on the IGF-IR and has been shown to be equivalent to or out perform insulin and IGF-I in supporting CHO cell growth and productivity.

 IGF-I, an animal-free recombinant supplement typically is used at 200-fold lower concentrations than insulin because of its increased IGF-IR affinity. It has a distinct biological advantage over other growth factor supplements because of a 100-fold reduced affinity for inhibitory IGF-binding proteins. Stimulation of the IGF-IR results in the activation of a number of signalling pathways, some of which are known to have key mitogenic and anti-apoptotic effects. LONG R

 IGF-I results in greater activation of these signalling molecules, thereby increasing culture longevity and productivity. LONG R

 IGF-I has been shown to be an effective alternative to insulin as a growth factor supplement for sustaining cell growth and viability in serum-free culture at industrial scale.

Transferrin is required to transport iron into cells, which is essential for cell growth and the regulation of key metabolic processes, such as DNA synthesis and oxygen transport.

  Transferrin has also been shown to play an important role in binding heavy metals in culture. Industrial cell lines such as CHO and NS0 require transferrin to attain optimal cell growth and productivity. Transferrin has been available in the form of serum-derived purified human transferrin (hTf) or bovine transferrin (bTf). Alternatively, inorganic iron salts have been used to supply iron to mammalian cells. To supply high-density cell cultures with sufficient iron, elevated concentrations of iron salts are required that use low affinity non-transferrin receptor pathways. This can have a negative effect on cell growth because of the formation of free radicals and oxidative stress from the unbound ferric or ferrous irons. Precipitation of iron hydroxide in the culture medium also can lead to limited bioavailability of iron to the cell.

Attempts to provide an efficient supply of iron by chemical chelators such as aurintricarboxylic acid or 2-hydroxy-2,4,6-cycloheptatrein-1-one (tropolone) has had limited application across a variety of cell lines because of unpredictability in controlling the intracellular redox cycle and cell oxidation processes.

A recombinant analogue of human transferrin, CellPrime rTransferrin AF (rTransferrin), (Novozymes) has shown equivalence to hTf and superiority to bTf in stimulating cell growth and productivity across a number of cell lines (Figure 1).

  rTransferrin binds specifically to the transferrin receptor, facilitating iron uptake into the cell. 

The demand for a universal and robust cell culture media has led to the need to identify cell culture components that substitute for the growth-promoting effects of serum. Nutrient requirements for individual cell lines have been found to differ considerably and, therefore, it has been difficult to design a single serum-free media for the growth of cell lines of commercial interest. In response to this demand, combinations of essential serum proteins have been examined for their ability to stimulate cell growth and productivity in a variety of industrially relevant cell lines. 

Studies in CHO cells have shown that growth and viability were adequately maintained in the absence of serum only when both IGF-I and transferrin were overexpressed in genetically engineered CHO cells.

  The combined action of two recombinant forms of these serum proteins, IGF-I (LONG R

 IGF-I) and transferrin (CellPrime rTransferrin AF) on CHO cell growth and productivity in SFM has been investigated (Figure 2). Results from this study show that a combination of these two recombinant proteins promote a synergistic increase in the levels of cell growth and productivity above those obtained from a standard SFM or each protein on its own.  

Biopharmaceutical companies require cell-culture media to be animal free, serum-free, defined, and cost-effective. In addition, there is a need to maintain or enhance process productivity while satisfying regulatory requirements for the elimination of serum components. A major driver for this has been the concern over contamination of the final drug product with adventitious agents derived from animal components. The industry has seen various attempts to produce robust animal-free, chemically defined media, and more recently, a protein-free media that is acceptable to regulatory agencies. However, the time involved in adapting cells to defined media, which often have resulted in reduced growth rates and product titers, has shown media development to be an important factor in the increasing cost of manufacture of the final drug product.

With a variety of recombinant, animal-free, defined protein supplements such as growth factors, transferrin, and albumin entering the market, the biopharmaceutical industry now has innovative and safer alternatives to serum and other animal-derived supplements. This situation allows manufacturers to rethink their media development to achieve greater process performance in a more regulatory compliant way.  

 is a senior scientist and scientific communications manager at Novozymes, Ltd., Thebarton, Australia, +61 883547787, 

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With a variety of recombinant, animal-free, defined protein supplements such as growth factors, transferrin, and albumin entering the market, the biopharmaceutical industry now has innovative and safer alternatives to serum and other animal-derived supplements.

The Role of Media Development in Process Optimization: An Historical Perspective

When expressing and purifying large quantities of soluble protein, expression difficulties often include poor yield and the formation of insoluble aggregates. Gene fusion technologies can overcome these obstacles and simplify purification and improve solubility. This article discusses the most popular fusion tags and the enzymes used to remove them, with special reference to recently introduced technologies.

Recombinant proteins show large variability in terms of their expression, solubility, stability, and functionality, making them difficult targets for large-scale analyses and production. Advances in recombinant protein expression include the development of better expression systems and host strains, improving mRNA stability, host-specific codon optimization, the use of secretory pathways, post-translational modification, co-expression with chaperones, and decreasing the amount of proteolytic degradation. However, no other technology has been as effective in improving the expression, solubility, and production of biologically active proteins as the addition of fusion tags, especially for difficult-to-express proteins.   

Genetically engineered fusion tags allow the purification of virtually any protein without any prior knowledge of its biochemical properties.

  They can improve the variable yield and poor solubility of many recombinant proteins. Proper design and judicious use of the right fusion tag can enhance the solubility and promote proper folding of the protein of interest, leading to recovery of more functional protein. On the other hand, adding fusion tags has been reported to result in changes in protein conformation, poor yields, loss or alteration of biological activity, and toxicity of the target protein. For this reason, it is desirable to remove the tag from the target protein after expression. When designing a fusion tag, therefore, careful consideration must be given to how the tag will be removed to produce native proteins without any extraneous sequences. Cleavage of the tags and the proteases that are used to cleave the tag are discussed in the next section. 

Many fusion tags are available for the expression and purification of proteins (Table 1). These tags can be broadly classified into two categories:

that aid in purification but do not enhance the solubility of the proteins substantially, and 

that specifically enhance the solubility and recovery of functional proteins.  

Table 1.  Commonly used fusion tags for purification and enhancing the solubility of proteins 

Affinity tags are the most commonly used tag for aiding in protein purification. They can be defined as exogenous amino acid (aa) sequences that bind with high affinity to a chemical ligand or an antibody. Most affinity tags are short peptide sequences that either bind to a ligand linked to a solid support (like the His tag) or contain an epitope recognized by immobilized antibodies (like the FLAG or Myc tags). The high affinity of these tags for their ligands and the availability of well developed immobilized supports for capturing the fusion proteins allow the protein of interest to be purified to a very high degree. Because of their small size, these affinity tags can be added at either end of the protein or in a region that is exposed to the surface. However, these tags generally do not increase the expression of the fusion proteins or enhance their solubility, and therefore are of little use in purifying hard-to-express proteins. 

His-tags are the most widely used affinity tags. The purification of his-tagged proteins is based on the use of a chelated metal ion as an affinity ligand;  one commonly used ion is the immobilized nickel-nitrilotriacetic acid chelate [Ni–NTA], which is bound by the imidazole side chain of histidine. Similarly, Streptag II, which consists of a streptavidin-recognizing octapeptide (WSHPQFEK), can be purified by affinity using a matrix with a modified streptavidin and eluted with a biotin analog. Other commonly used affinity tags like FLAG, Myc, and HA can be purified by binding to respective antibodies immobilized on chromatographic supports.  

Because it is desirable to remove most  tags at the end of the purification process,  considerable advances have been made in design of affinity tags so that they can be cleaved without leaving any residues behind and also to simplify the entire process of purification and cleavage. One such system is the "Profinity eXact" fusion-tag system (Bio-Rad, Hercules, CA), which uses an immobilized subtilisin protease to carry out affinity binding and tag cleavage. The protease is not only involved with the binding and recognition of the tag, but upon application of the elution buffer, it also serves to precisely cleave the tag from the fusion protein directly after the cleavage recognition sequence. This delivers a native, tag-free protein in a single step. Another system for simple purification of proteins is based on elastin-like polypeptides (ELP) and intein. ELP consist of several repeats of a peptide motif that undergo a reversible transition from soluble to insoluble upon temperature upshift. The fusion protein is purified by temperature-induced aggregation and separation by centrifugation, and intein is used for tag removal.

  No affinity columns are needed for initial purification. 

Solubility-enhancing tags are generally large peptides or proteins that increase the expression and solubility of fusion proteins. Fusion tags like GST and MBP also act as affinity tags and as a result, they are very popular for protein purification. Other fusion tags like NusA, thioredoxin (TRX), small ubiquitin-like modifier (SUMO), and ubiquitin (Ub), on the  other hand, require additional affinity tags for use in protein purification. 

No single fusion tag can increase the expression and solubility of all target proteins. However, some fusion tags have been more successful than others in increasing the solubility of many proteins. A comparison of some popular fusion tags showed that large proteins like NusA and MBP are more effective in solubilizing proteins than the smaller affinity tags or GST.

  Novel tags like Skip and T7 protein kinase, in turn, have been shown to be successful in expressing hard-to-express proteins in 

 Similarly, ubiquitin-based tags have been used to increase the solubility and expression level of proteins. SUMO tags are emerging as a viable alternative for increasing both the expression and solubility of otherwise hard-to-express proteins.

  The SUMO tag can be cleanly excised using SUMO protease, which recognizes the conformation of SUMO protein rather than a specific sequence within SUMO. Initially, the SUMO system was confined to 

 as highly conserved SUMO proteases are present in eukaryotes that cleaved the SUMO tag. However, the recently developed SUMOstar tag, a modified version of SUMO, is not recognized by the native eukaryotic protease and is specifically cleaved by the genetically engineered SUMOstar protease. Thus, the SUMOstar system can be used effectively in both prokaryotic and eukaryotic systems. The usefulness of the SUMO system was substantiated by the study of Marblestone, et al., who examined the effects of various fusion partners on total and soluble expression yield.

  They evaluated the expression and solubility of three model proteins fused to the C terminus of MBP, GST, TRX, NusA, Ub, and SUMO tags. The tags were ranked in terms of increased total expression as 

Overall, SUMO and NusA were equally good in terms of increasing the expression and the solubility of fusion proteins. However, SUMO offers certain advantages over NusA, in that SUMO is smaller and because it can be cleaved off precisely from the target protein without leaving behind any residues. 

The Rainbow tag is yet another new development in tag fusion technology. This technology allows the continuous monitoring of correctly folded proteins throughout the process of expression and purification. The Xavin mononucleotide (FMN)-binding domain of cytochrome P450 reductase (displaying a blue-green or yellow color, depending on the oxidation state of the FMN cofactor) and the red colored, heme-binding cytochrome b5 are used as tags, and the rainbow tags are visible with the naked eye.

  The use of rainbow tags, however, requires an additional affinity tag for purification. 

An integral part of the choice of a fusion  tag is the choice of the method for removing the tag after purification. This step almost always involves using a protease to cleave a specific peptide bond between the tag and the protein of interest. A small number of highly specific proteases are routinely used for this purpose and are listed in Table 2. These include the tobacco etch virus (TEV) protease;  thrombin (factor IIa, fIIa) and factor Xa (fXa) from the blood coagulation cascade; an enzyme involved in the cleavage or activation of trypsin in the mammalian intestinal tract, enterokinase (EK); proteases involved in the maturation and deconjugation of SUMO, SUMO proteases (Ulp1, Senp2, and SUMOstar); and a relative newcomer to the field, a mutated form of the 

 protease, subtilisin BPN' (Bio-Rad's Profinity eXact system). Many of these enzymes have been genetically engineered to enhance their stability (e.g., AcTEV, ProTEV) or their specificity, (e.g. SUMOstar, Profinity). With the exception of the SUMO proteases, all of these enzymes have the potential to cleave within the protein of interest.

  The SUMO proteases recognize not only their specific cleavage site, xaa-Gly-Gly/yaa, but also the tertiary structure of SUMO itself, giving them a very high degree of specificity. Bryan, et al., have  attempted to introduce the same level of specificity into the Profinity system by mutating both the subtilisin prodomain as well as the active site of subtilisin to increase the affinity of the enzyme for the prodomain and to decrease the likelihood of digestion within the protein of interest.

 One interesting consequence of this is that the affinity for the prodomain is so high that these researchers observed product inhibition of the enzyme. Essentially, the enzyme carries out one catalytic cycle and is then inhibited by the prodomain, which is retained in the active site, thus preventing further cleavage by this otherwise promiscuous enzyme. Because capture on the immobilized, mutant subtilisin matrix is an integral part of the system, the column must have a capacity (in moles of subtilisin) equimolar with the fusion protein. Although this is not problematic on the research scale, it could become prohibitively expensive at the multigram scale.  

Table 2. Proteases commonly used for tag removal 

The principle concerns with using a protease for removing a tag are 1) removing the protease following digestion, and 2) non-specific digestion of the target protein by the protease. Resolving the first concern is relatively straightforward, although in most cases it involves an additional chromatography step. Recombinant forms of TEV and its variants and of the SUMO proteases are all produced with a hexahistidine (His6) tag, allowing easy removal of the enzyme by metal chelate chromatography. Alternatively, some of these enzymes have been immobilized on solid supports, allowing their removal by simple filtration or centrifugation steps. Thrombin, fXa, and EK, which generally are produced from natural sources, can be removed by affinity chromatography, for instance, on benzamidine-agarose. With the Profinity system, cleavage and separation from the enzyme are combined in a single step. 

The second concern is more difficult to resolve. Non-specific cleavage is influenced by a number of parameters, such as the enzyme-to-substrate ratio (lower is better), temperature, pH, salt concentration, and length of exposure. TEV protease, thrombin, fXa, and EK all have well defined recognition sequences, but all of them have been found to cause "nicking" of the target protein in some instances. TEV protease has been re-engineered to try to increase its specificity (and stability), resulting in AcTEV (Invitrogen, Carlsbad, CA) and ProTEV (Promega, Madison, WI). Whether or not such engineering has reduced non-specific proteolysis remains to be seen. In addition, other tricks must be used with the native enzymes. For instance, one supplier recommends using fXa at pH 6.5, well below its pH optimum, to minimize non-specific cleavage. Of course, this requires the use of higher enzyme-to-substrate ratios and longer digestion times to achieve complete cleavage. Two of the enzymes listed (SUMO proteases and the Profinity enzyme) seem to be immune to this problem. SUMO proteases have evolved to recognize both the tertiary structure of SUMO as well as the cleavage sequence, xaa-Gly-Gly/yaa. The Profinity enzyme has been extensively mutated to derive a version that has very high affinity for the prodomain of the original enzyme. Thus, it also recognizes the tertiary structure of the prodomain as well as the cleavage sequence Phe-Met-Ala-Lys/yaa. On the other hand, SUMO proteases act catalytically (i.e., with a low enzyme-to-substrate ratio) whereas the Profinity enzyme requires equimolar concentrations of enzyme and substrate. 

One final consideration should be mentioned. Although one would ideally have a protein that is fully soluble in phosphate buffered saline at neutral pH, the reality is that for many proteins to be soluble at useful concentrations, they require more acidic or more basic pH levels, high or low salt levels, or the presence of chaotropes or detergents. It is therefore essential that the protease of choice retain substantial activity under adverse conditions. The most robust of the enzymes cited appear to be the SUMO proteases, the Profinity enzyme, and the TEV protease. Thrombin, fXa, and EK are much more sensitive to high salt concentrations or to the presence of chaotropes or reducing agents. 

Because every protein is unique, no single tag or cleavage method will answer every need. For proteins that express well, the simplest affinity tags may be sufficient (e.g., His6, myc). For harder to express proteins, fusion partners that enhance folding and solubility are preferable (e.g., MBP, SUMO). Tag removal then adds another layer of complexity. When considering which tag to use, key questions should be asked. For example, can your application tolerate retention of the tag, one or more amino acids remaining at the cleavage site, or must tag removal leave no trace? Such questions are usually answered experimentally, but with the availability of solubility-enhancing tags paired with highly specific proteases that cleanly remove the tag, these questions may be moot. 

 is vice president of research, all at LifeSensors, Inc., Malvern, PA, 610.644.6973, ext. 324, 

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 2. Esposito D, Chatterjee DK. Enhancement of soluble protein expression through the use of fusion tags. Curr Opin Biotechnol. 2006;17(4):353–8.  

 3. Banki MR, Feng L, Wood DW. Simple bioseparations using self-cleaving elastin-like polypeptide tags. Nat Methods. 2005;2(9):659–61. 

 4. Braun P, Hu Y, Shen B, Halleck A, Koundinya M, Harlow E, LaBaer J. Proteome-scale purification of human proteins from bacteria. Proc Natl Acad Sci. 2002;99(5):2654–9.  

 5. Hammarstrom M, Hellgren N, Van Den Berg S, Berglund H, Hard T. Rapid screening for improved solubility of small human proteins produced as fusion proteins in Escherichia coli. Protein Sci. 2002;11(2):313–21.  

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 7. Shih YP, Kung WM, Chen JC, Yeh CH, Wang AH, Wang TF. High-throughput screening of soluble recombinant proteins. Protein Sci. 2002; 11(7): 1714–19. 

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10. Marblestone JG, Edavettal SC, Lim Y, Lim, P, Zuo X, Butt TR. Comparison of SUMO fusion technology with traditional gene fusion systems: Enhanced expression and solubility with SUMO. Protein Sci. 2006;15(1):182–9.  

11. Finn RD, Kapelioukh I, Paine MJ. Rainbow tags: A visual tag system for recombinant protein expression and purification. Biotechniques. 2005;38(3):387–92.  

12. Jenny RJ, Mann KG, Lundblad RL. A critical review of the methods for cleavage of fusion proteins with thrombin and factor Xa. Protein Expr Purif. 2003;31(1):1–11.  

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Gene fusion tags can improve the yield and solubility of many recombinant proteins. This article discusses the most popular fusion tags and the proteases used to remove them, with special reference to recently introduced technologies.

Fusion Tags for Protein Expression and Purification

With the drive to remove serum and products of animal origin from cell-culture media during manufacture of protein-based biologics, plant hydrolysates have become popular as feeds in fed-batch cell culture to boost productivity. However, plant hydrolysates often contain undefined levels of nutrients, which can result in lot-to-lot variation in fed-batch performance and inefficiency. These are disadvantages when consistent protein yields are required from recombinant Chinese hamster ovary (CHO) cell lines. To achieve a more controlled process, one approach is to use chemically defined formulations as feeds containing only a carbon source, amino acids, vitamins, and trace elements. As a test of this approach, we compared cell growth and protein production characteristics of CHO cell lines grown in chemically defined media supplemented with either chemically defined nutrient feed supplements or plant hydrolysates alone, or with nutrient feed supplements.

Removing serum and products of animal origin from cell-culture media during production of therapeutic proteins from Chinese hamster ovary (CHO) cells offers improved biosafety and can make regulatory approval smoother. However, not having these supplements in the media can limit CHO cell growth and protein production. One method of improving growth is to add plant protein hydrolysates as a feed because they provide a highly pure source of soluble amino acids, peptides, vitamins, and essential elements for cell culture. Supplementing media with plant hydrolysates has been shown to improve protein production from engineered CHO cells.

The drawback of using plant hydrolysates is that they often contain undefined levels of nutrients and can result in lot-to-lot variation in fed-batch performance and purification inefficiency, which are disadvantages when consistent protein yields are required.

  To achieve a more controlled process, it is important to begin with a base medium for batch culture that does not make feeding strategies more complicated. An alternative approach to using nutrient-rich plant hydrolysates is to use a medium that is specifically designed to promote rapid cell growth and high levels of protein expression for CHO cells as a basal media, and then add chemically defined feeds containing only a carbon source, amino acids, vitamins, and trace elements to replenish the nutrients that are being depleted in the media.

  This article describes the experimental performance of chemically defined nutrient feed supplements compared to plant hydrolysates in a chemically defined medium with a recombinant CHO cell line. 

In a first shake flask study (Figure 1), recombinant CHO cells engineered to express a proprietary antibody were inoculated at 3 x 10

  cells/mL into 30-mL shake flasks containing CD OptiCHO Medium (Invitrogen, Grand Island, NY) as a base. The cells were incubated at 37 °C in 8% CO

, and agitated at 125 rpm. To determine the effect that plant hydrolysates or a chemically defined medium had on protein production, on days 3, 6, and 9, a 10% v/v solution of one of the plant hydrolysates derived from pea, wheat, soy, or rice (Kerry Bio-Science, Rochester, MN) was added at a stock concentration of 100 g/L to the cultures or the media were supplemented with a 10% v/v chemically defined feed, CHO CD EfficientFeed A (Invitrogen, Grand Island, NY). For each flask containing a different type of feed, there were three replicates made and samples were withdrawn for high performance liquid chromatography (HPLC) analysis of IgG concentration from each replicate every 24 hours on days 7–14. In those instances, when cell viabilities dropped below 20%, samples were no longer obtained. 

In a second shake flask study to determine if protein production could be increased by using a combination of feeds (Figure 2), 3 x 10

  cells/mL were inoculated into 30-mL shake flasks containing the same base medium. They were incubated at 37 °C in 8% CO

, and agitated at 125 rpm. On days 3, 6, and 9, a 10% v/v addition of the same solutions of pea, wheat, soy, or rice plant hydrolysates plus a 10% v/v solution of chemically defined feed A were added simultaneously, or a 10% v/v solution of chemically defined feed A was added alone to the cultures. For each flask containing a different feed combination, there were three replicates and samples were withdrawn for HPLC analysis of IgG concentration from each replicate every 24 hours on days 7–14. 

To determine the effects different feeds had on cell numbers and protein expression, a 15-L stir-tank bioreactor (Applikon Biotechnology, Foster City, CA) containing CD CHO medium was inoculated with a different proprietary recombinant IgG CHO cell line at 3 x 10

  cells/mL. The pH and dissolved oxygen were set at 6.9 and 50%, respectively, with agitation at 150–175 rpm. The CO

 and oxygen sparge rates were both set at 100 mL/minute. Glucose was added to 6 g/L if the concentration in the media in the bioreactor fell to less than 2 g/L. GS-Max (HyClone, Logan, UT) was fed at 1% of starting volume if the level of glutamate fell to less than 1.5 mM, as occurred in all bioreactors. Four different feed strategies were carried out over 10 days. In one bioreactor (Figure 3), 3% v/v addition of soy hydrolysate was added from a stock of 40 g/L every day for 10 days. In another bioreactor (Figure 4), a 3% v/v solution of chemically defined feed A, (CHO CD EfficientFeed A) was added on days 0–5 followed by addition on days 6–10 of daily supplementation of a 3% v/v solution of chemically defined feed B, (CHO CD EfficientFeed B). The additional two feed strategies determined the effects of higher concentrations of the chemically defined feeds but at fewer time points: 1) a 10% v/v solution of chemically defined feed A was added on days 0 and 3, followed by a 10% v/v solution of chemically defined feed B on days 6, 9, and 2) a 15% v/v solution of feed A was added on day 0 followed by addition of a 15% v/v solution of feed B on day 6. Both total cell counts and protein titers, as determined by ELISA over 13 days, are presented in the results (Figures 3 and 4). 

In the first shake flask study, addition of the plant hydrolysates produced protein yields of around 100–125 μg/mL (Figure 1). Culture viability using any of the hydrolysates did not extend beyond 12 days, while viabilities remained high with EfficientFeed A for the 14-day duration of the experiment. Interestingly, the soy and rice hydrolysates maintained viability for an extra day. This result is consistent with published data where the ability of soy hydrolysates to outperform other plant hydrolysates has been previously noted.

  The chemically defined feed attained protein yields of greater than 400 μg/mL, a four-fold yield improvement compared to any of the plant hydrolysates tested. 

In the second shake flask study, addition of the plant hydrolysates and the chemically defined feed produced protein yields from around 175–375 μg/mL (Figure 2). Addition of chemically defined feed and plant hydrolysates together extended cell viability to 14 days with all plant hydrolysate combinations and resulted in increased protein yields by as much as three-fold when compared to the performance of plant hydrolysate alone. Soy and rice hydrolysate combinations with the chemically defined supplement generated the highest yield, while pea and wheat hydrolysate combined with chemically defined feed generated the lowest bioproductivity. However, when the chemically defined feed was added alone, it achieved higher protein yield than the feed and plant hydrolysate combinations. The lower yields obtained by adding plant hydrolysates and chemically defined feed together may be explained by osmolality increases, although this was not investigated, and therefore, cannot be proved in this study.  

In the bioreactor study, after the daily addition of 3% v/v soy hydrolysates, a total cell count of 7 x 10

  cells/mL was achieved at day 9, compared to 9–11 x 10

  cells/mL with various combinations of defined supplement and feeding strategies (Figures 3 and 4). The addition of soy hydrolysate resulted in a lower cell growth rate up to day 9, when the soy bioreactor was terminated because of contamination. The growth rates of cells grown in the presence of 10% and 3% v/v chemically defined feeds were similar and both achieved a maximum cell count of ~11 x 10

  cells at day 9, while those grown in 15% v/v chemically defined feeds were lower, with cells achieving a maximum cell density of ~9 x 10

Cell density does not necessarily have a direct relationship to protein expression. Cells supplemented with 15% feed A on day 0 and an addition of 15% feed B on day 6, which achieved the highest cell density, expressed comparable protein production values to those grown with other feed strategies. Cells grown in the presence of 3% v/v soy hydrolysate compared to the 3% v/v chemically defined feeds showed slightly less than half the cell density, yet protein production was only around 10% lower (0.7 mg/mL at day 9 with soy hydrolysate and 0.8 mg/mL at day 9 with chemically defined medium). Lower osmolality for the soy hydrolysate condition may have contributed to the lower productivity seen. Overall, the best bioproductivity resulted when using a counter-intuitive feeding strategy—feeding with a relatively large bolus of supplement, both at initiation of culture and at one more timepoint before the culture lag phase. Therefore, consideration of multiple defined nutrient supplement formulations is suggested when maximum protein expression is required. 

Results demonstrated that for the recombinant CHO cell line studied, the addition of chemically defined feed to the basal medium produced approximately a four-fold greater protein yield than any of the four different plant hydrolysates used. Also, culture supplementation with different plant hydrolysates in combination with chemically defined feeds provided no advantage in terms of increased protein production over the addition of a chemically defined feed on its own. Supplementing media with plant hydrolysates may not always be the most efficient choice or may not offer the same increases in productivity available from chemically defined feeds. Therefore, when culturing CHO cells for maximum protein production, it may be worthwhile to compare different feed types. Bioreactor studies of a different CHO cell line also demonstrated that chemically defined feeds in the culture provided slightly higher protein yields than the addition of a soy hydrolysate when the two feeds were added at the same concentration. In addition, using a higher concentration of chemically defined feed A on day 0 followed by feed B on day 6 yielded superior bioproductivity and proved the value of including multiple, and sometimes counter-intuitive, feeding strategies when trying to maximize protein production. 

In summary, using chemically defined feeds with CHO cell lines not only eliminates the variability associated with using plant hydrolysates, but could also improve the productivity of biopharmaceutical protein manufacture and help move therapeutic proteins into clinical trials more rapidly. 

 is a technical area manager of media development, 

 is a principal scientist, Cell Systems Division, 

 is a senior product manager, all at Invitrogen Corporation, Grand Island, NY, 716.774.6860, 

 1. Mols J, Peeters-Joris C, Wattiez R, Agathos SN, Schneider YJ. Recombinant interferon-gamma secreted by Chinese hamster ovary-320 cells cultivated in suspension in protein-free media is protected against extracellular proteolysis by the expression of natural protease inhibitors and by the addition of plant protein hydrolysates to the culture medium. In Vitro Cell Dev Biol Anim. 2005. Mar–Apr; 41(3–4):83–91.

 2. Burteau CC, Verhoeye FR, Mols JF, Ballez JS, Agathos SN, Schneider YJ. Fortification of a protein-free cell culture medium with plant peptones improves cultivation and productivity of an interferon-gamma-producing CHO cell line. In Vitro Cell Dev Biol Anim. 2003 Jul–Aug;39(7):291–6.  

 3.  Luo Y, Chen G. Combined approach of NMR and chemometrics for screening peptones used in the cell culture medium for the production of a recombinant therapeutic protein. Biotechnol Bioeng. 2007 Aug 15;97(6):1654–9. 

 4.  Stein A. Decreasing variability in your cell culture. Biotechniques. 2007 Aug;43(2):228–9.  

 5.  Chun BH, Kim JH, Lee HJ, Chung N. Usability of size-excluded fractions of soy protein hydrolysates for growth and viability of Chinese hamster ovary cells in protein-free suspension culture. Bioresour Technol. 2007 Mar;98(5):1000–5.

Using chemically defined feeds with CHO cell lines not only eliminates the variability associated with using plant hydrolysates, but could also improve the productivity of biopharmaceutical protein manufacture and help move therapeutic proteins into clinical trials more rapidly.

Improving Protein Production in CHO Cells

Recombinant protein and plasmid DNA production using microbial expression systems is the cornerstone of many biologics manufacturing processes. High cell density (HCD) methods are commonly used for these processes because of the advantages they provide, including high cell productivity, high concentration levels, and lower setup costs. Additionally, the increased yield in product translates to a more cost-effective and shorter overall project length.

As the fields of DNA vaccines and gene therapy mature, many companies are using high cell density production methods that have proven to be effective in the production of recombinant proteins. However, adopting a high cell density process for DNA production has some unique challenges. This article will examine some of the considerations that should be evaluated before adopting high cell density fermentation for DNA production. Specifically, which type of production (batch, fed-batch, or continuous fermentation), which media and components, and which strategies for growth control for the high cell density methods will be discussed.  

The production costs associated with developing biopharmaceuticals is an increasingly important consideration for companies that develop these products. As products approach commercialization and companies have to reconcile the high cost of production with relatively low reimbursement rates, this consideration receives even more attention. There are many aspects of production that can be evaluated and optimized. Specifically, production costs may be reduced through efficient vector design, strain selection, and optimizing upstream and downstream production processes. Significant efforts have been made to increase the productivity of recombinant proteins produced in 

, including the use of high cell density fermentation processes. The progression of plasmid DNA products in development pipelines has also prompted the application of high cell density processes in the production of 

High cell density (HCD) processes have many inherent advantages. Specifically, HCD reduces the time required in a fermenter in either a contract manufacturing facility or in captive space. Secondly, plasmid processes can produce yields as high as 1–2 g/L, which is a marked improvement over standard fermentation methods, and this results in fewer required fermentation runs. Various HCD process parameters have been evaluated in the development of a large-scale process for the production of clinical-and commercial-grade plasmid DNA. 

The first consideration in designing an efficient process should be the vector design and host cell line selection. Plasmid size is a critical criteria in vector design. All nonessential sequences should be removed so that the plasmids are as small as possible. In addition to the potential regulatory and therapeutic challenges, many larger plasmids also create manufacturing hurdles by placing a metabolic burden on the host cell line by reducing resources required for plasmid replication. This, in turn, results in reduced yields.  

Formulation is also a consideration that must be made at the stage of vector design. Although naked DNA has inherent advantages over more complex formulations, including safety and simplicity, transfection efficiencies are generally low. There are strategies that may be used during vector design, including translational engineering, that can remove secondary structure and add translational pause sites, which may increase expression. 

The use of ampicillin and other B-lactams is discouraged because of hypersensitivity reactions in some patients.

  If they are used, the FDA often requires a justification and details of the precautions that should be taken to prevent these reactions.

  Kanamycin is the most commonly used means of selection. Cell line selection is one of the most critical criteria in the design of HCD processes. Bacterial cell lines offer an advantage over mammalian cell lines in that many cell lines may be evaluated simultaneously and very quickly for either protein or plasmid production. In addition to evaluating yields, the purity and quality of the plasmid in various cell lines will impact manufacturing decisions. In our experiments, some cell lines, such as DH10β, and to a lesser extent, DH5α, have been found to be consistently higher producing in HCD processes.

Once the vector has been designed and the host cell line selected, one of the first decisions that must be made is whether to initiate production with batch, fed-batch, or continuous fermentation processes. With the demands for plasmid DNA in late clinical trials approaching the 100-gram scale or greater , a batch fermentation, with typical yields of 10–20 mg/L, becomes unfeasible. Limitations are caused by uncontrolled growth rates and waste product accumulation in this mode. A better choice for avoiding these issues and thereby increasing productivity is the use of fed-batch or continuous high cell density fermentation. Continuous fermentation processes are conducive to the production of large amounts of a single product but maintaining sterility is a major challenge. Fed-batch fermentation, which starts with a short batch fermentation followed by the addition of media at a defined rate, offers many advantages for DNA production, particularly in circumstances where many vector types are produced. Fed-batch offers more flexibility and consistency than batch production. Specifically, it allows for relatively simple optimization of fermentation profiles for each plasmid DNA product. Even with modifications to fermentation profiles from plasmid to plasmid, this feed strategy produces controlled cell growth results that are inherently more consistent and predictable than batch fermentation. Scale-up from process development activities to production of commercial quantities is a fairly streamlined process.  

The most straightforward feed strategy is a defined growth rate strategy. When using a defined growth rate, feed media is added at rates as determined by a pre-established growth profile. In our process, the feed is triggered by a DO

 spike, which is caused by the exhaustion of the initial bolus of glucose in the media. Following this initial spike, the program feeds media based on a predefined feed profile. 

Media composition can dramatically affect yields and consequently the overall cost of production. Options vary greatly, including minimal (defined) media to complex (semi-defined). Complex media formulations often contain ingredients like yeast extract, peptones, and other growth factors that may allow for a higher cell density, but may present challenges with reproducibility and with contaminant removal in downstream processing. Minimal media contains known quantities of essential nutritional components including a carbon source, a nitrogen source, and salts, and excludes components known to be inhibitory to bacterial growth. Fermentation processes using minimal media are highly reproducible and plasmid copy number may even be higher when using minimal media.

  Our process uses a minimal media for its high cell density manufacturing processes. Minimal media also allows for adjustment and optimization through the addition of components that may increase yields. Glucose is routinely used in 

 fermentations and was chosen as the carbon source. It is easily obtained in a purified form and easy for the 

Recombinant protein and plasmid DNA production using microbial expression systems is the cornerstone of many biologics manufacturing processes. HCD methods are commonly used for these processes because of the advantages they provide.

BioPharm International-06-02-2008

Recombinant Protein Production Yields from Mammalian Cells: Past, Present, and Future

The Role of Media Development in Process Optimization: An Historical Perspective

Fusion Tags for Protein Expression and Purification

Improving Protein Production in CHO Cells

Maximizing Yields of Plasmid DNA Processes