Development and optimization of growth media, feed solutions, and feed strategies is a complex project. More than 200 components (amino acids, trace metals, vitamins, growth factors, carbon sources, etc.) can be found in various commercial growth media formulations. A typical cell culture medium formulation may contain 50 to 60 individual components. Some of these may be critical for cell growth or productivity, others may be toxic at certain levels, and many may be involved in complex interactions in the same or competing pathways within the cell. Media and feed components typically are screened individually or in small combinations in shake flasks, in experiments of a few dozen shake flasks in parallel. This method significantly limits the ability to discover complex interactions among medium components. 

A high-throughput medium development technology that allows the rapid development of optimized base growth media, feed solutions, and feed strategies has been developed and was used in the following study with a proprietary Chinese hamster ovary (CHO) cell line. Approximately 100 components were screened in titration experiments to select the top 16 components for feed optimization. A 16-component central composite design with axial and center points was then implemented in a shaker plate model system. Based on the productivity levels, two of the top combinations were selected for confirmation in shake flask studies. Using a high-throughput approach, we were able to increase the viable cell density and volumetric productivity from 3.5 x 10

/mL and 1.2 g/L in two experiments. This article discusses the high-throughput technology and the experimental approach.  

A recombinant CHO cell line engineered to express a proprietary antibody was provided by an outside party. Cells were grown in a commercial serum-free medium with L-glutamine and insulin. Cell cultures were grown at 37°C with 5% CO

. Initially, optimal conditions were determined for the growth of CHO cells in a shaker plate model system, and it was shown that cell growth was comparable to shake flasks. A high-throughput liquid handler was used to deliver the samples to the shaker plates according to the combinations derived from a design of experiments software program. For this study, 23 amino acids and derivatives, 4 carbon sources, 10 hydrolysates, 8 lipid solutions, 21 salts, 7 trace metals, 18 vitamin solutions, and 9 other components were prepared in the serum-free medium (without L-glutamine or insulin); pH was adjusted to 7.2 ± 0.2. 

Cell count and the viability of cultures in flasks were determined using trypan blue exclusion dye. Viable cell counts for the cultures in the shaker plate system were determined using a high-throughput plate-based imaging method in which a fluorescent viability stain was added, and plates were incubated and scanned using a high-throughput fluorescent microscope. Protein concentration was determined using a bead-based enzyme-linked immunosorbent assay (ELISA)-style immuno-competition assay. Streptavadin beads were prepared with a biotinylated anti-immunoglobulin G (IgG) antibody. Unknowns were competed against a Cy5-labeled IgG and read against an antibody standard. 

For feed formulation development, titration experiments were performed with 100 components, from which 16 components were selected based on their impact on viable cell days, specific productivity, and prior experience. A 16-component central composite design with triplicates was performed in the shaker plate system. Shake flask studies were performed using the top two combinations based on protein productivity from the 16-component design.  

Figure 1. High throughput media development strategy			

Dilution series of selected feed components were performed to check for optimal concentration ranges and toxicity levels. These experiments were performed in a shaker plate system. A serial dilution was performed from 2x through 0x. The plates were incubated at 37°C, 5% CO

, and were run for 7 to 9 days. Cells were sampled on days 3, 5, 7, and 9 for cell viability assays. For one set of experiments, it took one day to set up the experiment; after incubating the plates for 7 to 9 days, significant components were selected based on cell viability and protein productivity assays. A schematic of the media development strategy is shown in Figure 1.  

Table 1. Combinations from the 16-component central composite design			

Approximately 16 components were selected for feed optimization based on the cell growth curves from feed formulation experiments. A 16-component central composite design was set up in triplicates in a shaker plate system. Cells were seeded at a density of 1.85 x 10

/mL, and the plates were incubated at 37°C, 5% CO

, and were run for 7 to 9 days. Cells were sampled on days 3, 5, 7, and 9 for high-throughput cell viability and protein productivity assays. Two of the top combinations from the shaker plate system were selected for shake flask studies based on protein concentration. The factorial design of some of the combinations is shown in Table 1.  

Figure 2. Cell viabilities of the control, run 258, and run 179 in triplicates are shown from day 0 to day 9. The controls maintained good viability for 5 days and samples 258 and 179 maintained good viability for 7 to 8 days.			

Shake flask studies were performed with combinations 179 and 258, as shown in Table 1, to confirm the results from the shaker plate system. These studies were performed in triplicates along with the controls. The feed components were added according to the combinations, and the controls were added with the same amount of media but no additional components. This process was followed to ensure the same volumes in all the experimental shake flasks. The initial seeding density after adding the components was 1.72 x 10

/mL. Cells were sampled on each day up to 9 days. Cell viability and cell counts are shown in Figures 2 and 3. The controls reached a maximum viable cell density of 4 x 10

Figure 3. Viable cell counts of the control, run 258, and run 179 in triplicates are shown from day 0 to day 9. The maximum viable cell count for the controls was on day 5 and for runs 179 and 258 was on day 6 and day 7, respectively.			

Viable cell density for run 258 was maximal on day 7, with greater than 7 x 10

/mL on day 9. Run 179 had the highest viable cell density, reaching 8 x 10

/mL by day 9. The average protein productivity of the cells with two different feed combinations performed in shake flasks is shown in Figures 4 and 5. Figure 4 shows the average protein concentration at the time of harvest, and Figure 5 shows the daily cumulative average protein concentration. At viable cell densities correlating with cell harvest conditions, the average protein concentration was 0.3 g/L for the control, 0.9 g/L for run 258, and 1.2 g/L for run 179. The protein productivity of samples 258 and 179 was three to four times greater than the control.  

Figure 4. The figure shown here is the average protein concentration of triplicate samples in g/L. Control and runs 179 and 258 were compared when culture viability dropped below 50%. The error bars indicate Â±1 standard deviation.			

Growth of CHO cell cultures in a shaker plate model system was shown to be comparable to growth in shake flasks. This model system enabled the use of automated liquid handling and plate-based assay techniques to be combined with design of experiments to create a high-throughput platform for medium and feed development. To develop an optimized feed formulation, 100 cell culture medium components were screened and tested in 16-factor central composite design experiments. The results of the top two feed formulations were confirmed in shake flasks to increase productivity three to four times, up to 1.2 g/L. Each experiment took one day to set up and up to nine days to execute; thus, a media development program of this scope can be conducted by one researcher in approximately one month.  

Figure 5. Cumulative average protein productivity of the triplicate control samples and the 179 and 258 run triplicate samples on day 3, day 5, day 6, day 8, and day 9.			

 is a senior research associate at Xcellerex, Inc., Marlborough, MA, 508.480.9235, 

 is vice president of process development and technology, all at Xcellerex Inc., Marlborough, MA. 

Articles

Growth of CHO cell cultures in a shaker plate model system was demonstrated to be comparable to growth in shake flasks.

Rapid High-Throughput Feed Optimization

Over the past decade, disposable technologies have become a reality in biotech processes. The use of disposables in research and manufacturing allows high flexibility. 

The aim of cultivation in a bioreactor is to produce biomolecules using animal or plant cells, microorganisms, yeast, and insect cells. 

Biomolecules are divided into four main classes: carbohydrates, lipids, nucleic acids and proteins.

  The latter is the most common in biotech processes: manufacturing of vaccines, diagnostics, or therapeutic preparations.  

Mammalian cells constitute a demanding system for the production of heterologous proteins. Two main formats have been employed for the production of recombinant proteins in mammalian cells: cultures of adherent cells and suspension cultures. The latter is by far the most common.

  In 2004, mammalian cell-based therapeutic proteins reached a market share of 59% followed by 27% for 

Before production can start, the right cell line is screened out and the cultivation process must be optimized, for example, scaled-up, starting cultivation with a few milliliters and ending up with several cubic metres' cultivation volume. 

For more than 40 years, typical bioreactors for cell cultivation have been made of glass or stainless steel and have been used in research and commercial production processes. This traditional bioreactor was characterized by substantial investment cost resulting from the necessity for aseptic bioprocess technology, sterilization in place (SIP), cleaning in place (CIP) and validation, all requiring sophisticated instrumentation. 

Disposable bioreactors with mechanical energy input represent modern alternatives to such traditional cultivation systems for some applications in lab- and pilot-scale. 

The main characteristics of the single-use bioreactor are low cost, ease of operation, time saving and high process security. These parameters shorten the time-to-market. 

Disposable bioreactors can be divided into two groups: 					

Bag reactors with mechanical energy input where the cells are directly in contact with growth media. The most common bag reactor is based on rocking motion, where the wave-induced motion guarantees the energy input.

Membrane cultivation systems with two chambers separated by a semipermeable membrane. The membrane enables the passage of nutrients to the cells, yet the metabolites can leave the cell chamber. High cell densities can be reached, but up-scale capability is limited to the low lab-scale level.

This article aims to outline the potential and limits of the disposable bioreactor based on wave-induced agitation for biotechnology use, while considering the working principle.  			

This disposable bioreactor with wave agitation can be used for batch, fed-batch and a perfusion operation mode. It is shown that even fragile cells such as animal cells (CHO, NS0, fibroblasts and hybridoma) human cells (T-Lymphocytes, HEK and Perc.6), insect cells (Sf9, High5 with baculovirus) and plant cells (hairy root culture, suspension cultures and embryo culture) can be grown in sealed bags made of polyethylene.

Disposable bioreactors based on the rocking motion principle are mechanically-driven reactor systems, which are available in lab- (up to 25 L) and pilot-scale (up to 300 L) versions. They consist of three components: 					

A rocker base unit containing a bag holder with heating capabilities.

The biocompatible disposable chamber with integrated, single use, sensors for pH and DO.

The measuring and control unit (Figure 1).

Figure 1. BIOSTAT CultiBag RM optical.										

Energy input is affected by rocking the chamber back and forth, generating a fluid movement in the cell culture and medium (Figure 2). In this way, the surface of the medium is continuously renewed so bubble-free aeration can take place. Therefore, a maximum of half of the total bag volume can be filled with the cultivation media. The other half is needed for the overlay aeration. Depending on the sensitivity of the cultivated cell line, the angle and amplitude of the rocking motion, as well as velocity can be varied. Specially designed sterile plastic cultivation bags contain tubing for filling, harvesting and sampling. This allows simple handling and optimal cell growth for different cell types and cell lines.  

Figure 2. Principal of rocking mixing technology.			

The measuring and control unit can facilitate regulation of pH, DO, temperature, rocking rate and gas flow using the gas mixing system. An example for gas supply strategy is shown in Figure 3. The pO2 cascade control and pH control with CO2 additionally to base and acid is provided.  

Figure 3. Aeration strategy with BIOSTAT RM Tower optical (Sartorius BBI Systems).			

The used cultivation bags can be sterilized by autoclaving and then destroyed by incineration. The PE foil from which the cultivation bag is made, is environmentally compatible when incinerated. This is less critical for the environment than the cleaning of conventional bioreactors with alcohol, acids and basic solutions. An additional safety factor is the fact that no cell or protein residues are released into the wastewater cycle. 

The advantages of disposable bioreactors are mainly ease of use, prevention of cross-contaminations, and cost and capital investment savings. The costs compared with conventional stainless steel bioreactors are lower because the capital investment requirement is lower, and the necessity for qualification and validation processes, as well as maintenance, is reduced. A cost reduction of up to 41% compared with conventional stirred bioreactor technology is possible.

  Additionally, the disposable bioreactors enhance process safety because the contamination risk falls under 1%.10 The disadvantage of using disposable bioreactors is the limitation in scale-up (300 L) and aeration capabilities for cultivation of some microorganisms, which require high oxygen uptake rates. 

Oxygenation or aeration can be achieved by three methods, individually or in combination; that is, membrane aeration, surface aeration or direct sparge aeration.  

Microbial fermentations exhibit high metabolic oxygen demand together with high biomass concentration; this combination requires an energy-intensive means of maintaining adequate culture aeration. This aeration is achieved by simultaneously sparging air at high rates and vigorously agitating the culture to increase the bubble interfacial area and affect the mass transfer of oxygen. 

The biomass concentrations and cellular oxygen uptake rates observed in mammalian cell cultures are low compared with microbial systems (Table 1). Even if the cellular oxygen uptake rate is low, it does not mean that oxygenation does not present a problem. The reason is the shear sensitivity of mammalian cells, which prevents the use of vigorous aeration or agitation. As a result, the volumetric oxygen mass-transfer coefficient is low and becomes a limiting factor in scale-up.  

Table 1. KLa(h-1) and OTR(h-1) values of different cultivation systems (source: Enfors 2003 and Wave Biotech AG, Switzerland).			

Bioreactors for cultivating mammalian cells are typically aerobic bioreactors and oxygen is usually a limiting nutrient because of its low solubility in culture media. When bioreactors are scaled up from laboratory to production size, their design must meet both oxygen distribution and oxygen mass transfer requirements. 

One engineering parameter that could be used to compare the newly developed disposable bioreactors with established stainless steel cell-culture bioreactors could be the oxygen transfer rate (OTR). Because of its low solubility, only 0.3 mM O2, equivalent to 9 mg/L, dissolves in one litre of water at 20 °C in an air/water mixture. This amount of oxygen will be depleted in a few seconds by an active and concentrated growing culture unless oxygen is supplied continuously. In contrast, during the same period the amount of other nutrients used is negligible compared with the bulk of concentrations. Therefore, most aerobic processes are oxygen-limited. This is the reason why the concept of gas–liquid mass transfer in bioprocesses is centered on oxygen transfer even if other gasses such as carbon dioxide, hydrogen, methane and ammonia can also be involved.   			

The mass transfer of oxygen into liquid can be characterized by the OTR or by the volumetric oxygen transfer coefficient (KLa). The larger the KLa, the higher the aeration capacity of the system. These values have been thoroughly examined as a critical parameter for bioreactor function.   			

Disposable bioreactors with wave agitation are used for commercial and R&D applications for the production of cell-derived products using animal, insect, plant and virus cultures in good manufacturing practice (GMP) and non-GMP applications. Because of their flexibility, single-use cultivation bags are especially suitable for screening new cell lines, media optimization and vaccine production. Basically, two strategies can be implemented when considering the use of disposable bioreactors:  					

A partially disposable strategy; for example, preparation of inoculum in disposable cultivation bags for process-scale cultivation in conventional stirred tank reactors (Figure 4).

Figure 4. Disposable system approach.					

A completely disposable strategy, where inoculum as well as plant bioreactors are disposables based on the rocking-motion principle (Figure 5).

Figure 5. Disposable system approach.										

In addition, the use of multiple large-scale disposable bioreactors instead of one conventional stirred tank reactor can be considered, particularly in view of their ease-of-use and process safety against contaminations of disposable bioreactors. Finally, the parameter that will document the merits of using conventional bioreactors or disposable bioreactor systems for biotechnology processes are yield of production, expenditure of time and process costs.

  The cultivation of yeast and aerobic microorganisms is, however, a challenge in disposable bioreactors. As shown in Table 2, because of the published Kla or OTR values, it is unlikely that a wave agitated system can be used for the most common microbial expression systems because of the Kla or OTR values inherent in this method.  

Table 2. Oxygen uptake rate values for mammalian cells and E. coli.			

Even if the market for disposable bioreactors has grown rapidly in recent years, full acceptance will only be reached when the entire production process from media preparation, inoculum preparation and downstream processes can be done using disposable equipment. Some suppliers are focusing on this issue to become a total solution provider for disposable biotech factories. 

 is a product manager of disposable bioreactors at Sartorius AG, Switzerland. 

is a scientist in process development, biotechnology, at Sartorius AG, Germany. 

 1. L. Stryer L. Biochemistry, 4th ed. New York (NY): W.H. Freeman and Company; 1995. 

 2. Wurm F. Nature Biotech 2004;22: 13–98. 

 3. Sandig V. et al. Mammalian cells. In: Gelissen G., editor. Production of recombinant proteins. (Wiley-VCH; 2005)  

 4. Weber W. et al. Catching the Wave: The BEVS and the Biowave. Novartis Pharma Research, Switzerland. 

 5. Eibl D. and Eibl R. BioWorld 2005; 3:1–2 (Supplement BioteCHnet)  

Energy input is affected by rocking the chamber back and forth, generating a fluid movement in the cell culture and medium.

Disposable Bioreactors Based on Wave Agitation Technology

The US Food and Drug Administration (FDA) and the Department of Agriculture have erected a series of barriers to protect humans from exposure to the fatal agent linked to bovine spongiform encephalopathy (BSE), and measures announced by the FDA in January 2007 build on those barriers. With its heavy reliance on mammalian cell-based expression systems, and with the problem of serum-free versus protein-free versus productivity, the biopharmaceutical industry continues to need chemically undefined or defined supplements of animal and human origin. To move toward avoidance of animal- or human-derived materials used during biopharmaceutical manufacturing, the industry is increasingly looking at alternative raw materials and is investigating the benefits of non-mammalian expression systems that offer simpler and chemically defined growth conditions. New recombinant raw ingredients are becoming more readily available and have been developed specifically for use in large-scale mammalian cell culture. Animal-free microbial expression systems have made such products possible, and these may themselves offer solutions to some regulatory and productivity issues the biopharmaceutical industry faces.  

Biotherapeutics—including recombinant proteins, monoclonal antibodies, and nucleic-acid-based drugs—represent the largest growth sector in the pharmaceutical industry, with a market size estimated at $33 billion and projected to reach $70 billion by the end of the decade. Of the 31 therapeutic proteins approved since 2003, 17 are manufactured using mammalian cell lines.

  Mammalian cell culture is the method of choice for manufacturing large complex proteins, especially those requiring posttranslational modifications, such as glycosylation.  

Traditionally, a range of animal-derived protein products—including fetal calf serum, peptones derived from acid or enzyme, hydrolysates of casein, gelatin, meat, egg, lactalbumin, and bovine serum albumin (BSA) or human serum albumin (HSA)—have been used for mammalian cell culture media for research and commercial-scale manufacturing of biopharmaceuticals. These ingredients, together with supplements such as human holo transferrin and insulin-like growth factors (IGF), are added to promote cell growth, survival, and optimized productivity. 

REGULATORY REQUIREMENTS FOR THE PRODUCTION OF PHARMACEUTICALS

Due to the threat of contamination from bovine materials infected with BSE and the threat of transmission of variant Creutzfeldt-Jakob disease (vCJD) from HSA, regulatory authorities worldwide have issued guidelines that govern the use of animal-derived materials in pharmaceutical products. For example, both the European Medicines Agency (EMEA)

  and the Center for Biologics Evaluation and Research (CBER) of the US Food and Drug Administration (FDA)

  have issued guidelines for the controlled use of animal-derived materials for pharmaceuticals from sources with a risk of BSE or vCJD infection.  

In addition to existing guidance, in January 2007 the FDA announced further proposals to prohibit the use of certain bovine materials as ingredients in some medical products or as elements of product manufacturing. These proposals are the latest in a series of BSE safeguards that would bar material that harbors the highest concentrations of this fatal agent in infected cattle.  

To ensure that companies comply with these prohibitions, the FDA proposes that records be kept to demonstrate that any bovine material used as an ingredient in these medical products or as part of their manufacturing process should meet FDA requirements. This proposal is in addition to measures that companies must already take to reduce risks from serum-derived products, including: 					

testing for the presence of adventitious agents

inactivation or removal treatments for contaminating agents.

Current efforts in the US to minimize the risk of BSE infection from pharmaceuticals are in line with those in place in the EU since 2003.

  These US regulations require manufacturers of medicinal products to undertake and document extensive risk assessments on all animal-derived materials used in any aspect of pharmaceutical production, including in the preparation of active substances, excipients, adjuvants, and raw starting and packaging materials. Those assessments should be  based on: 					

the nature of animal-derived material used and cross-contamination procedures

systems in place for ensuring product consistency and traceability.

Today the use of animal-derived materials in pharmaceutical production places an increasingly onerous burden on product manufacturers to comply with established requirements. 

ADVANCES IN RECOMBINANT PROTEIN ALTERNATIVES

With the construction of further barriers to the use of animal-derived materials, there are growing needs and trends in the industry to move toward the use of serum-free, chemically defined, and most recently, protein-free media formulations for use in mammalian cell culture. Using chemically defined media components not only reduces the risk of introducing adventitious agents, but it also assists both upstream and downstream process optimization. 

While major advances have been made in this area, there are still drawbacks to the use of these products, especially for industrial-scale culture.

  Transfer to serum-free or protein-free media can be very stressful for cells, and it typically requires a period of adaptation or "weaning." Successful adaptation can be time-consuming and cell-line-dependent, and a lack of protein components can lead to a less robust and less reproducible process. 

It is recognized that the addition of specific growth factors and proteins (which are naturally found in serum) can enhance cell viability and productivity. Serum-free media, which is still used extensively in research laboratories, can contain proteins that may or may not be animal- or human-derived. Similarly, to enrich protein-free media, hydrolysates may be added that are not always non-animal in origin.  

Recently, dedicated research and development into the production of defined, recombinant alternatives to three critical components of serum (insulin, transferrin, and albumin) have led to a range of animal-free recombinant protein ingredients that are designed and manufactured specifically to optimize mammalian cell growth and productivity at the industrial scale. With these ingredients, researchers should now be able to achieve the performance associated with protein-supplemented media while avoiding the associated risks of traditional animal-derived supplements.  

Studies with recombinant ingredients have shown equivalent or improved performance in a wide range of cell types up to commercial scale.

  Manufactured to recognized quality standards and offering continuity of supply, these ingredients provide the confidence in critical raw material quality that is required of products used in long-term biopharmaceutical manufacturing. 

Growth factors are essential for long-term growth and proliferation of cell lines in serum-free media formulations. Recombinant therapeutic insulin has been used as a growth factor in serum-free cell culture. However, its primary use for the treatment of diabetes has led to supply and availability issues for cell culture users.  

Recombinant insulin-like growth factors are now available as a dedicated raw material, manufactured exclusively for the industrial cell culture market. Studies using this mitogen as a supplement in cell culture have shown that it results in performance that is equivalent to or better than recombinant insulin in numerous cell types, including: Chinese hamster ovary (CHO), baby hamster kidney (BHK), human embryo kidney-293 (HEK293), Vero, PER.C6, Madin-Darby canine kidney (MDCK), and fibroblasts (data not shown).  

 IGF-I is a recombinant insulin-like growth factor (IGF-I) analogue manufactured in 

 Studies comparing the effectiveness of LONGR

 IGF-I with insulin for sustaining cell growth and viability in serum-free CHO cell cultures have found that recombinant LONGR

 IGF-I protein is better able to sustain viability under production conditions than is insulin.

  Insulin is mostly used at supra-physiological concentrations, and its ability to promote growth and survival is attributable to its activation of the IGF-I receptor at these high concentrations. LONGR

 IGF-I is also more potent than insulin at activating the IGF-I receptor, even at 200-fold lower concentrations in both CHO and HEK293 cells (Figure 1).

Figure 1. Combined results from subculture experiments showing the percentage cell growth with DeltaFerrin-supplemented serum-free media relative to growth obtained with holo human transferrin and other commercially available chemical chelators (ferrous ammonium sulphate, ferrous sulphate, and Invitrogen B), over five subcultures in Vero-PP, Madin-Darby canine kidney (MDCK), and baby hamster kidney (BHK)-21-PP1-C16			

Animal- and human-derived transferrins have traditionally been used as cell culture supplements to facilitate optimal iron metabolism. Studies in CHO cells have found that a combination of both IGF-I and transferrin is necessary to maintain viability and proliferation upon the withdrawal of serum from the culture media.

In an effort to move away from animal-derived transferrins, investigators look to using combinations of chemical-based iron chelators. However, a recent study questioned the ability of the chelators to consistently support a wide range of cell lines in serum-free media. Additionally, in comparing various commercially available iron chelators (ferrous ammonium sulphate, ferrous sulphate, and Invitrogen B) against a plasma-derived  recombinant human transferrin (DeltaFerrin), the authors found that the recombinant human transferrin (DeltaFerrin) was the only alternative growth factor that showed almost equal potency to native human holo transferrin over five subcultures in Vero-PP, MDCK, and BHK-21-PP1-C16 (Figure 2).

Figure 2. Combined results from subculture experiments showing the percentage cell growth with DeltaFerrin-supplemented serum-free media relative to growth obtained with holo human transferrin and other commercially available chemical chelators (ferrous ammonium sulphate, ferrous sulphate, and Invitrogen B), over five subcultures in Vero-PP, Madin-Darby canine kidney (MDCK), and baby hamster kidney (BHK)-21-PP1-C16			

The use of chemical chelators for industrial-scale manufacturing may also be limited by the chelators' unpredictability in controlling and managing the redox cycle and cell oxidation processes. An animal-free recombinant human holo transferrin, specifically designed and manufactured for industrial cell culture applications, provides the benefits of natural plasma-derived transferrin, with the defined, regulatory-friendly aspects of chemical alternatives.  

Albumin is a 66-kDa plasma protein with a variety of physiological and stabilizing functions in vivo.

  In cell culture, albumin (usually in the form of BSA or HSA) is known to serve several important functions, including acting as a carrier protein and protecting cells from mechanical stress, and it is also thought to be a major serum survival factor, functioning as an inhibitor of apoptosis.

Several recombinant human albumin (rHA) products are now commercially available and are offered as alternatives to HSA for a range of biopharmaceutical applications. rHA expressed from 

 has been shown to exhibit similar safety, tolerability, and pharmacokinetic/pharmacodynamic profiles to native human albumin,

 and it has shown equivalent capacity to protect immunological, biological, and biochemical cell function.

 (for example, rProbuminAF), is being developed specifically for industrial cell culture use. Used as an ingredient in mammalian cell culture, such recombinant albumins can deliver quality, regulatory benefits, and performance benefits for both research and commercial-scale applications.  

ANIMAL-FREE RECOMBINANT SUPPLEMENTS—A STEP IN THE RIGHT DIRECTION 

New animal-free cell culture ingredients derived from microbial expression systems (such as those described above) offer a valuable solution to the quality and performance limitations of supplement regimes currently used with serum-free and protein-free media formulations for industrial-scale mammalian cell culture. Such ingredients, designed and manufactured exclusively to support biopharmaceutical manufacturing, bypass many regulatory hurdles and offer a secure, consistent supply for commercial-scale production.  

ANIMAL-FREE TECHNOLOGY: THE MICROBIAL OPTION

Mammalian expression systems historically have  been characterized as slow and expensive vehicles for the large-scale manufacturing of therapeutic proteins. They are, however, essential for the production of therapeutics that require complex post-translational modifications (PTMs), particularly glycosylation. O-linked and N-linked glycosylation patterns can influence protein stability, ligand binding, immunogenicity, and serum half-life, and they are significant in the context of efficacy of a wide range of biopharmaceuticals.

On the other hand, microbial systems as models for the animal-free production of recombinant proteins on an industrial scale have much to offer the biopharmaceutical industry. 

Unlike mammalian-based technology, microbial systems are typically grown in simple, chemically defined media. And, in the case of the proprietary 

systems (which enabled the development of the recombinant ingredients described above) the entire production process is free from the use of animal- or human-derived materials.  

With faster growth rates, high yields, and well understood genetics, microbial systems remain the obvious choice for expression of non-glycosylated peptides and proteins. Of the 31 therapeutic proteins approved since 2003, nine are produced in 

   However, in spite of this prokaryote lacking the ability to perform eukaryote -specific PTMs, it is widely used for the production of insulin, growth hormones, and growth factors. Yeast-based systems confer the advantages of high expression levels, easy molecular manipulation, and low costs of goods; in addition, as eukaryotes, yeast can perform PTMs, such as proteolytic processing, folding, disulphide bond formation, and glycosylation. 

Since the 1980s, yeasts have been used for the large-scale production of recombinant proteins of human, animal, and plant origin. The brewer's yeast 

 is an extremely well characterized eukaryotic system with "generally regarded as safe" (GRAS) status, and it was widely used for early protein production, including the first commercialized recombinant vaccine.

 has recently suffered from a negative image in the industry due to a number of perceived disadvantages in comparison to methylotrophic yeasts (e.g., 

). These perceived disadvantages include  mitotic instability of recombinant strains, undesirable over-glycosylation, and difficulties in adapting production strains to industrial scale.

 systems offer strong promoters, stable integration of expression plasmids, and high-density fermentation. However, they also have a number of limitations at large scale, including the use of hazardous chemicals (e.g., methanol induction), lack of moderately expressed promoters, and few selectable markers.

Advances in the molecular biology and process development of 

have overcome many of the issues outlined above, and have produced an effective microbial expression technology for the animal-free production of certain classes of biotherapeutics.

  Extensive strain development has created a proprietary 

-based system, delivering a highly competitive cost of goods, typical of a microbial fermentation-based process.  

 strains now exist that typically grow to high cell densities in short fermentation cycles in animal-component–free chemically defined media. Various desirable traits—including genetic stability, high copy number expression plasmids, protease deficient mutants, and strains deficient in the enzymes involved in O-linked glycosylation—have been engineered.

  The systems are optimized for the production of recombinant proteins where glycosylation does not naturally occur or can be designed out without impacting product efficacy. This yeast-based system also overcomes many problems associated with prokaryotic alternatives, such as improper disulfide bond assignment, risks of protease degradation, and inclusion body formation. 

The constitutive expression system has not given rise to any problems with post-translational modifications, such as proteolysis or other forms of degradation that would make a separate induction phase advantageous. To produce recombinant proteins in high yields and of optimum quality, 

 has been subjected to a series of genetic manipulations. The success of the design of the disintegration vector, designed as whole 2-μm vectors in an otherwise plasmid-free background, dispels the common view that yeast episomal plasmids are unstable for industrial use, as no plasmid loss occurs during the production process in semi-continuous operation over a time scale of months.  

Commercial-scale manufacturing of Recombumin—the first recombinant human albumin approved by both the EMEA and the FDA for use in the production of human therapeutics—has successfully validated the disintegration vector system and the automatic procedures developed in the laboratory for the control of the fed batch process function at plant scale.  

In addition to existing guidance, in January 2007 the FDA announced further proposals to prohibit the use of certain bovine materials as ingredients in some medical products or as elements of product manufacturing.

Advances in Animal-Free Manufacturing of Biopharmaceuticals

Rapid, efficient, and cost-effective protein expression and purification strategies are required for high throughput structural genomics and the production of therapeutic proteins. Fusion protein technology represents one strategy to achieve these goals. Fusion protein technology can facilitate purification, enhance protein expression and solubility, chaperone proper folding, reduce protein degradation, and in some cases, generate protein with a native N-terminus. No technology or reagent is a panacea, however, and establishing tools and optimal conditions for each protein remains an empirical exercise. With this in mind, protein fusions are a leading option to produce difficult-to-express proteins, especially in 

 a simple and low cost host, has long been the preferred system for recombinant protein expression. Recently however, difficult-to-express proteins (e.g., G-protein coupled receptors [GPCRs], kinases, ion channels, blood plasma proteins, vaccines, and antibodies) have become the norm instead of the exception. This trend is steering researchers away from 

 to more costly higher organisms such as baculoviruses and mammalian cells.  

The recombinant therapeutic protein business had sales of more than $34 billion in 2004.

  This total excludes therapeutic antibodies, vaccines, DNA–RNA synthetics, small molecules, and gene and cell therapies. Projections to 2010 show sales amounting to $52 billion. Table 1 lists top selling recombinant therapeutic proteins.   

Table 1. Top eight recombinant therapeutic proteins and their global sales between 2002 and 2004.			

In the future, the industry is likely to be hampered by an imposing and growing bottleneck resulting from an inability to efficiently express large quantities of biologically active protein at low costs. The so-called "low hanging fruits" have been picked and a new era involving difficult-to-express proteins is upon us. This situation may drive the industry to use more costly higher organisms for protein production.  

The application of gene fusion technology to 

 however, offers a promising alternative. Commercial success is more likely when the expression host is simple and inexpensive.  

Typical problems with expressing difficult-to-express proteins include: low or null expression yields; insoluble protein; purification difficulties; degradation of the expressed protein; and incorrect folding. Some advances in improving recombinant protein expression in 

 include the development of strong promoters,

  and most influential of all, fusion tags. Examples of popular fusion tags include: glutathione-s-transferase (GST),

Gene fusion technology can facilitate purification, enhance protein expression and solubility, chaperone proper folding, reduce protein degradation, and in some cases, generate protein with a native N-terminus. Nevertheless, protein expression remains an arduous task that involves a complex decision tree. Whether or not to use gene fusion technology is just one choice. Other factors include the expression system, host strain, mRNA stability, codon bias, inclusion body formation and prevention, site-specific proteolysis, secretion, post-translational modification, and co-overexpression. The complexity is compounded by the diversity of proteins. To date, no technology or reagent is a panacea. Thus, establishing tools and optimal conditions for each protein remains an empirical exercise. 

Recently, several comparative studies have examined the effects of various fusion partners on total and soluble expression yield (Table 2). Marblestone et al. evaluated the expression and solubility of three model proteins fused to the C termini of MBP, GST, TRX, NusA, Ub, and SUMO tags.

				 The tags were ranked in terms of increased total expression as TRX > SUMO ~ NusA > Ub ~ MBP ~ GST and increased soluble expression as SUMO ~ NusA > Ub ~ GST ~ MBP ~ TRX. Hammarstrom et al. cloned 27 human proteins (MW < 20 kDa) into various expression vectors and ranked the tags' ability to promote soluble expression as TRX ~ MBP ~ Gb1 > ZZ > NusA > GST > His6.

  Braun et al. compared the expression of 32 human proteins (molecular weight of which varied from 17 to 110 kDa) and ranked tags in terms of increased expression and yield after purification as GST ~ MBP > CBP > HIS6.

  Shih et al., in a study of 40 different proteins with eight different tags, observed that MBP gave the best overall results in terms of total and soluble expression.

  In one of the studies in Dyson et al., the solubility of 20 mammalian proteins was compared and the fusion tags were ranked in terms of increased soluble expression as TRX ~ MBP > HIS10 > GST > GFP.20 De Marco et al. demonstrated that NusA was better than GST at enhancing the solubility and stability of recombinant proteins.

Table 2. Summary of several comparative studies that examine the effects of various fusion partners on total and soluble expression yield.			

The inconsistency of the data from these comparative studies only solidifies the statement that tools and optimal conditions for each protein remain empirical and that no technology or reagent is a panacea. Nevertheless, it is likely that in the future generalities about specific fusion tags may be made (e.g., for entire protein classes). As the comparative studies suggest, gene fusion tag systems range dramatically in efficiency. 

Factors that influence yield and biological activity include: a) the affinity purification scheme; b) enhancement of recombinant protein expression; c) protein folding and enhanced solubility; d) protection from degradation; e) size of the fusion tag; and  f) the specificity, efficiency, and site of cleavage. Herein lies a further discussion of these factors. 

Protein fusion tags are often used to purify proteins from crude extracts. GST and MBP are two examples. Other protein fusion systems, such as NusA, SUMO, Split SUMO, thioredoxin, and ubiquitin require an affinity tag, such as polyhistidine (HIS). Numerous examples of affinity purification exist for fusion proteins, including nickel-nitriloacetic acid to isolate hexahistidine-fused proteins,

  and GSH-sepharose to isolate glutathione (Table 3).

  Successful purification schemes achieve high quality and quantity with inexpensive, high capacity resins and mild elution conditions. For effective purification, high affinity between the fusion tag and resin is essential, but affinity must not be too high, because harsh elution conditions can disrupt tertiary structure.  

Table 3. Affinity tags influence protein expression yield and activity.			

Enhancing Recombinant Protein Expression 

Protein expression depends on transcriptional regulation, mRNA stability, and translational efficiency, whereas enhanced recombinant protein expression is governed by a high mRNA copy number, efficient translational initiation and elongation, stability of the mRNA, and the translational enhancers (reviewed by Makrides).

  Codon bias is another factor that affects expression,

  yet it has been overcome by engineering new strains or cell lines that contain rare tRNAs or by altering the problematic codons to more common prokaryotic codons.

Promoters also play a fundamental role in the transcription of heterologous genes and recombinant protein expression. Strong and highly regulated promoters are now commonplace for 

  On the other hand, there is still much to be learned about gene fusion technology, which has been shown to dramatically enhance expression.

  The exact mechanism by which fusion proteins enhance expression remains unknown. Some speculate that it is the result of the highly conserved structure of the fusion tag.

Protein Folding and Enhanced Solubility  

Cost and simplicity are the primary driving forces when choosing a recombinant expression organism. As a result, 

 has various shortcomings as a recombinant expression organism. Many eukaryotic proteins, especially proteins with disulfide bridges or sugar moieties, cannot be expressed as soluble, active, and properly folded proteins in 

 often yields macromolecular crowding (200–300 mg/mL in the cytoplasm), which presents an unfavorable environment for protein folding and results in a high concentration of incorrectly folded proteins that form undesirable inclusion bodies that require re-folding. Inclusion bodies afford protection from proteolytic degradation, which may be their only advantage. 

To circumvent these problems, several strategies have been implemented to enhance solubility, promote properly folded protein, and reduce the percentage of inclusion bodies. These strategies include co-expression of molecular chaperones and foldases, expression of secreted proteins, and expression of protein fusions.

Protein fusion tags have been shown to act as solubility enhancers and chaperones.

  Neither mechanism is well understood, but the hypotheses include: 					

Fusion of a stable or conserved structure to an insoluble recombinant protein may serve to stabilize and promote proper folding of the recombinant protein.

Fusion tags may act as a nucleus of folding ("molten globule hypothesis").

It should be noted that even though fusion partners promote solubility, this is not a universal indicator of correct folding, and researchers recommend taking additional measurements (including monodispersity by light scattering,

  CD spectropolarimetry, bis-ANS binding,

  ligand binding or enzymatic activity) to provide supporting evidence for correct folding. 

Recombinant proteins often are considered unwanted by cells and are subjected to proteolytic degradation.

  Several strategies have been developed to protect recombinant proteins from degradation, including the use of protease inhibitors,

  secretion into the periplasm or culture medium,

  The compartmentalization hypothesis describes the mechanism by which gene fusions protect against proteolytic degradation.

  Fusions can promote the translocation of their partner proteins to different cellular compartments, thereby decreasing the concentration of the recombinant protein in the protease-rich cytosol. For example, SUMO can translocate from the cytosol to the nucleus and maltose binding protein (MBP) can translocate to the membrane compartment of the cell.

The size of the fusion tag plays a crucial role in the overall yield of the purified protein. As seen in Table 4, the yield of the purified target protein is dictated by the yield of the fusion protein: the larger the fusion tag, the lesser the overall yield. Split SUMO, which is 47 amino acids long, is only 19% of the fusion when Split SUMO is fused to a target protein that is 200 amino acids in length (47/247). In contrast, NusA, which is 495 amino acids long, is 71% of the fusion with the same target protein (495/695). Therefore, if expression yielded one gram of fusion protein for both Split SUMO–Target and NusA–Target, the yield of the purified target after cleavage would be 0.810 g and 0.288 g, for Split SUMO and NusA, respectively.  

Table 4. The size of the fusion tag influences the yield of the purified protein.			

Protein expression remains an arduous task that involves a complex decision tree. Establishing tools and optimal conditions for each protein remains an empirical exercise.

BioPharm International-05-02-2007

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