				Perfusion processes in mammalian cell cultivations are currently experiencing a renaissance, and are making their way into modern single-use bioreactor systems. In this article, the authors review the common challenges and main applications for perfusion processes. The authors present experimental evidence for reaching extreme cell densities of 150 ×106 cells/mL in an easy to use, high-performance, single- use bioreactor that provides an efficient way to generate cell mass for cell banking or seed trains, as well as production of considerable amounts of monoclonal antibodies (mAbs), even from cell lines with mediocre expression levels.

During the past decade, the pharmaceutical industry has increasingly switched focus from small molecule-based drugs to recombinant therapeutic proteins, mAbs, vaccines and more recently, cell therapy products to find novel treatments for unmet medical needs. Considerable effort has been dedicated to improving the efficiency of cell lines, vectors, culture media and the production process itself. Today, a number of platform approaches to high-titer expression, especially of mAbs, have been reported (1, 2). These platforms consist of a host cell line, vectors, media, and feeds that are crucial to reach titers of >2–5 g/L in a 14 day fed-batch process. 

MAIN DRIVERS FOR USING  PERFUSION CULTURE 

Recently, there has been renewed interest in perfusion processes. In perfusion culture, fresh medium is continuously fed to the culture while the cells are retained in the bioreactor, and metabolic waste products are removed. Depending on the chosen cell retention device, the large molecular product can either be retained in the bioreactor (concentrated fed-batch) or removed from the culture (perfusion). The latter production mode is typically used when the recombinant protein is prone to degradation, toxic, or inhibitory to the cells. In perfusion processes, cell densities are typically 5– to 10–fold higher than in batch or fed-batch processes. Process optimization of perfusion processes is intended to maintain cell-specific productivity by optimizing the composition of the perfused medium and/or applying a certain cell bleed stream to control cell density and growth rate. It is then possible to achieve 5 to 10 times higher total amount of product compared with batch and fed-batch (3). An economic comparison of the different process modes has recently been published (4). Due to significantly increased productivity, a number of companies have identified concentrated fed-batch and perfusion as tools to reduce production scale and ultimately make commercial production amenable to single use bioreactors. This growing interest in continuous perfusion is further fueled by the increased number of recombinant therapeutic proteins in the development pipeline.  

Several cell-retention devices have been developed during the past 20 years. Today, there are a handful of systems that have proven suitable for suspension cell culture at process scale: continuous centrifugation, inclined settlers, hydrocyclones, and alternating tangential filtration (3, 4). 

In addition to improved yields in protein production, high cell-density culture also offers advantages for seed production by reducing the bioreactor scale and number of steps necessary to generate the seed for the final production bioreactor. A 10–fold increase in cell density reduces the required inoculum volume by the same factor. To increase flexibility and reduce the time from thawing a cell bank vial to inoculation of the production bioreactor, studies have been performed examining the use of large volume, cryopreserved cultures in cryobags, where the concentrated seed culture has been produced using a perfusion process in order to replace shake flask technology (5, 6). This approach not only reduces the number of bioreactors and steps involved, but also the necessary footprint for seed bioreactors, and investment in equipment.  

CHALLENGES WITH  CONVENTIONAL CELL-RETENTION SYSTEMS

Perfusion often requires expensive investment in cell-retention equipment, certain infrastructure, and personnel with specialised skills. Therefore, it has not been widely adopted for the production of proteins that are used for research purposes, such as target proteins or therapeutic lead proteins used in early preclinical research. At the same time, early protein supply is often characterised by low expression rates of the cell lines used and the need to produce many different proteins in a given period of time to fuel the development pipeline. We have therefore developed a novel, easy to use, benchtop perfusion bioreactor that is based on rocking motion agitation (BIOSTAT CultiBag RM 20 perfusion, Sartorius Stedim Biotech). In this study, we present data on the high cell-density culture of a mAb-producing Per.C6 model cell line. 

CELL PROPAGATION AND BIOREACTOR INOCULATION

A Per.C6 cell line expressing a monoclonal anti–EpCAM (epithelial cell adhesion molecule, a cancer marker) immunoglobulin G (IgG) antibody (licensed from Crucell) and commercially available, chemically-defined serum-free medium, CDM4 PerMab (Hyclone), was used for the batch and perfusion phases of the cultivation. Inoculum was produced in two 1 L polycarbonate Erlenmeyer flasks (Corning) with 300 mL culture, each incubated in a Certomat CT plus shaker incubator (Sartorius Stedim Biotech) at 85% humidity, 37 °C, 5% CO

The bioreactor bag (CultiBag RM10 perfusion, Sartorius Stedim Biotech), equipped with single-use optical pH and DO probes, was filled with 5 L of medium and equilibrated to process temperature. After calibration of the optical sensors, the reactor was inoculated with a seeding density of 5 x 10

  cells/mL. The initial process parameters were 19 rocks/min, a rocking angle of 6° and a temperature of 37 °C. The pH was controlled at pH 7.1 with CO

. Dissolved oxygen (DO) was controlled at 40% by automatically adjusting the gas flow rate (range 0.3 to 3 L/min) and the composition of N

. For comparison to the perfusion process, a batch culture was performed using the above mentioned parameters.  

An empty bag was welded to the perfusion outlet of the bioreactor bag, and a 50 L media bag was welded to the perfusion inlet and placed on a balance. Perfusion was started at an exchange rate of 0.5 reactor volumes/d, which corresponds to a feed and harvest flow of 0.105 kg/h. The perfusion rate was subsequently increased based on the measured glucose and glutamine concentration as detailed below. As the bioreactor controller automatically increased the gas flow rate and oxygen content in the process gas, the angle and rocking rate were adjusted manually from initially 6° to 10°, and 19 rocks/min to 23 rocks/min. Due to the increased angle and rocking rate, oxygen transfer from the headspace to the liquid phase could be further improved. 

				Samples were taken at regular intervals for offline pH measurement (Sartorius PB-11 pH meter) and monitored for cell count (Nucleocounter, Sartorius Stedim Biotech), glucose, glutamine, and lactate concentration (Bioprofile 100plus Analyzer, Nova Biomedical) and antibody concentration (ELISA).			

HIGH CELL DENSITY CULTIVATION OF AN ANTIBODY EXPRESSING CELL LINE

In batch mode, a cell density of 17 × 10

  cells/mL at a viability of 98% was reached (see Figure 1). This result is typical for a pH and DO controlled bioreactor batch culture of this specific Per.C6 clone and the PerMAB medium used. Once the substrate was depleted, cells went into the stationary phase. In contrast, in an uncontrolled process, e.g., in a shaker flask, the cell density typically reaches about 12 × 10

  cells/mL under similar conditions (data not shown). 

Figure 1: Viable cell density and viability of batch and perfusion process. 			

In perfusion culture using the BIOSTAT CultiBag RM 20, we were able to reach a very high cell density of 150 × 10

  cells/mL at a viability of 99%. Up to now, the highest reported cell density of a wave-type, single-use bioreactor has been 60 × 10

  cells/mL (7). Typically, we achieve about a 10-fold higher cell density in perfusion culture using this bioreactor set up compared to the batch results (data not shown). The total cell yield was 7.5 × 10

  cells derived from 5 L of reactor culture. 

It was possible to obtain very high cell densities without the need for sophisticated equipment, fed-batch strategies or process optimization by using off-the-shelf equipment with powerful control capabilities. The L–glutamine and glucose concentrations were used to initiate the perfusion mode and to determine the medium exchange rate. When the glutamine concentration dropped below 1 g/L, which was 66 h after inoculation, addition of fresh medium started. In order to maintain the glutamine concentration at above 0.5 g/L, the perfusion rate was increased step-wise. After 138 hours, the perfusion rate was increased to 1.5/d (0.315 kg/h), and after 163 hours, to an exchange rate of 3/d (0.63 kg/h) (see Figure 2). 

Figure 2: Glucose, glutamine and lactate concentrations over the course of the culture. 			

The perfusion process was terminated after 9 days when the perfusion membrane started to block; the perfusion control system detected the reduced harvest flow rate and shut down the pumps. Consequentially, the remaining substrate was consumed, lactate accumulated, and the logarithmic growth phase ended. Typically, in a production setting, one would directly move to harvest of the supernatant. In our set up, we wanted to test the limit and robustness of the bioreactor system. In the case of cell production, e.g., for inoculation of a production bioreactor or large scale cell banking in cryobags, the culture would be terminated before reaching the limit of the perfusion membrane. 

 The signals from the disposable optical DO and pH sensors that are integrated into the single use bioreactor bag were used to control the respective process parameters. pH was controlled mainly by adjustment of CO

 in the gas stream. Disposable optical sensor technology is a rather new, but versatile, technology which is suitable even for challenging applications like this high cell density perfusion culture. Regular off-line measurement using a conventional pH meter served to check the precision of the optical pH sensor, and to determine if recalibration of the optical pH sensor would be necessary to compensate for a potential drift. Also, changes in ionic strength might affect the accuracy of the sensor readout. During culture, the ionic strength might be influenced by accumulation of lactate or ammonia or other metabolic by-products. A comparison of the pH values obtained from the disposable optical sensors and the off-line values is shown in Figure 3. At the beginning of the perfusion process, one recalibration of the optical pH sensor based on the off-line value was performed. During the course of the high density culture, pH could be maintained at the set point despite a change of the perfusion rate, which temporarily led to an increased pH in the bioreactor. 

Figure 3: Biorector pH control and comparison between online and offline pH data. 			

Process parameters affecting DO in a rocking motion bioreactor are shown in Figure 4. The DO could be maintained throughout the whole high cell density culture at approximately 40%. The rocking rate was increased manually from 19 rocks/min to 21 rocks/min, and finally to 23 rocks/min. The angle was increased from 6° to 7° and finally to 10°. As the rocking rate and angle increased, the wave formation in the bag became stronger, hence increasing the surface exchange rate at the gas-liquid interface and ultimately the oxygen transfer rate (OTR) of the bioreactor. The OTR could further be increased by adding pure oxygen to the process gas. Since we worked with moderate rocking rates, which leave room for further increase for most common cell lines, it can be concluded that we are still far away from the upper limit of the oxygen transfer capacity that can be achieved in this type of bioreactors. It should be noted that the increase of the rocking angle from 7° to 10° at process time 160 h reduced the requirement of pure oxygen in the process gas dramatically. 

Figure 4: Dissolved oxygen concentration (DO), gas flow, percentage of pure oxygen (O2) in gas stream, rocking rate and angle. 			

INCREASE IN PRODUCTIVITY AND CELL DENSITY 

We have compared the productivity of recombinant anti–EpCAM in the bioreactor system operating in batch and perfusion mode. During the perfusion run, approximately 50 L of medium had been consumed, which was 10 times the amount that was used in batch mode. The maximum viable cell concentration was increased by a factor of 10.  The total IgG yield in the perfusion run was 5900 mg, which was approximately 30 times greater than in the batch mode, where the total amount was 205 mg (see Figure 5). This illustrates that the perfusion system is perfectly suited for the fast production of high amounts of proteins such as monoclonal antibodies. 

Figure 5: Total Immunoglobulin (IgG) amount in batch and perfusion cultivations. 			

The authors demonstrated that an easy to use, off-the-shelf, single-use, bench-top perfusion bioreactor is capable of producing and supporting very high cell density cultures of up to 150 × 10

  cells/mL. This approach can be applied to mediocre cell lines in order to quickly produce the amounts of recombinant proteins typically required during drug discovery or early development (e.g., for high throughput screening, or early preclinical studies in animals), and formulation development. Furthermore, this is a simple approach to improve the efficiency of seed production in large scale production. A typical seed train for a 1000 L bioreactor consists of a 50 L and 200 L bioreactor applying a dilution factor of 1:5 to 1:10 at maximum. Assuming an inoculum cell density of 5 × 10

  cells/mL at 1000 L scale, which is equivalent to 5 × 10

  cells, this density can be achieved by approximately 20  L of a high cell density culture grown in a rocking-motion perfusion bioreactor. Certain limitations of this technology could be expected in larger bag systems, such as 100 L or 200 L rocking motion bags with integrated membranes for cell retention. The oxygen transfer rate in these systems is usually lower, and the ratio of membrane surface area to culture volume can not be scaled in a linear fashion due to size restrictions. While cell densities of 30-40 × 10

  cells/mL have been reported for these larger scale rocking motion systems, it is likely that extreme high cell densities as reported in this article may not be reached. 

* is product manager, single use bioreactors, 

 is head of process engineering, upstream technology, 

 is vice-president, marketing and product management, fermentation, all at Sartoius Stedim Biotech, Göttingen, Germany, 

 3. C. Fenge and E. Lüllau, "Cell Culture Bioreactors" in 

Cell Culture Technology For Pharmaceutical and Cell-Based Therapies

, S. Ozturk, and W-S Hu, Eds. (Taylor & Francis, New York 2006), pp. 155–224. 

 6. A. Amanullah et. al., Cell Culture Engineering IX conference, (Cancún, Mexico, March 2004). 

Articles

Peer-Reviewed Research

An off-the-shelf, single-use perfusion system.

Increasing Efficiency in Protein and Cell Production by Combining Single-Use Bioreactor Technology and Perfusion

Drugs, biological therapeutics, and vaccines manufactured under cGMP must be controlled at all stages of the production process. The final release testing is not in itself sufficient to guarantee quality, safety, purity and efficacy of the final product (1). It is necessary to ensure that the manufacturing operations perform as intended within the established specifications. This implies that any given process is well characterized, adequately controlled, and reproducible. Fermentations are complex processes used in the manufacture of recombinant proteins and peptides, vaccines, and other therapeutic products. The performance of fermentation (and cell culture) processes are critical to the quality and consistency of production of the therapeutic products and are influenced by parameters, such as pH, temperature, dissolved oxygen, as well as nutrients and other components necessary for cell growth, and metabolites. The nutrients include carbohydrates, amino acids, and other organic compounds, which act as carbon sources essential for cell growth and product synthesis. Inorganic cations (metal ions) also play critical roles for optimum activities of enzymes and in cell growth. The metabolic products sometimes act to inhibit the product synthesis and affect the desired yield. Thus, measurements of nutrients and metabolites in fermentation broths (and culture media) are important for understanding the roles they play and microbial physiology, as well as for the development, optimization, characterization, and control of the process. Such measurements also permit improvement of the process performance by providing the ability to determine the need for supplementation, as well as to the development of defined media to replace complex media formulations. Most of the nutrients and metabolites, including inorganic ions, carbohydrates, alditols, alcohols, and aliphatic carboxylic and amino acids are ionic or polar in nature, and do not have chromophores necessary for analysis by absorption measurements. Ion chromatography (IC) with electrochemical detection (ED) presents a suitable technique for the determination of these components. The technique usually requires no sample preparation other than dilution with water. Other chromatographic methods often require complex pre- or postcolumn derivatization for the detection of analytes. 

This article will provide a brief overview of IC and ED, as well as their applications in the analysis of nutrients and metabolites in fermentation broths and neomycin in selective culture media. 

IC is a form of high-performance liquid chromatography (HPLC), which involves separation based on ionic interactions among ionic or polar analytes, ions present in the mobile phase (eluent), and ionic functional groups on chromatographic support (stationary phase). This can lead to two distinct mechanisms of separation based on the type of interactions—(a) ion exchange in which the analyte ions are adsorbed to and desorbed from the stationary phases repeatedly leading to separation, and (b) ion exclusion due to repulsion between similarly charged analyte ions and the ions on the chromatographic support. Separation based on ion exchange has been the predominant application of IC to-date.  

In ion-exchange chromatography, ionic functional groups on the chromatographic support have charges opposite to those of the analyte ions. That is, a column for separation of cations, a cation-exchange column, contains negative ions. This technique is widely used in the analysis of cations, including metal ions, anions, mono- and oligosaccharides, alditols (sugar alcohols), aminoglycosides (antibiotics), amino acids, peptides, organic acids, amines, alcohols, glycols, thiols, and other polar molecules. Ion-exclusion chromatography uses strong cation- or anion-exchange columns to separate ionic, polar, weakly polar, and apolar analytes. The ionic analytes experience strong repulsion from similarly charged functional groups on the chromatographic support and are eluted first, with little or no separation. An apolar analyte experiences no repulsion and is, therefore, eluted late from the column. Polar analytes experience less repulsion than an ion but more than an apolar molecule and are separated in order of their polarity (more polar compound eluting earlier). This technique has been used in the analysis of organic acids, alcohols, glycols, and sugars.   

Any suitable detector can be used for the detection and quantitation of analytes by IC. However, two forms of ED, suppressed conductivity detection (SCD) and pulsed amperometry detection (PAD), have distinct advantages over other detection methods. They provide the selectivity and sensitivity necessary for the measurement of critical components of fermentation broths without any pre- or post-column derivatization or complex sample preparation procedures.  

When a constant voltage is applied across two electrodes between which the effluent from a column flows, a current is generated because the effluent contains ions and polar molecules. The current is proportional to the conductivity of the solution, which, in turn, is proportional to the concentration of ionic species in solution, leading to the detection and quantitation of polar analytes by a conductivity detector. The problem, however, is that the conductivity of the mobile phase is significantly higher than the conductivities of the analytes, simply because the concentrations of ions in the mobile phase is 104–105 times higher than that of the analytes. SCD permits detection and quantitation of analytes by reducing the conductivity of the mobile phase (baseline conductivity) to nearly zero. It also enhances the conductivity of the analytes, thereby enhancing sensitivity by about an order of magnitude (2). The peaks of apolar and weakly polar molecules, such as sugars (except the ionic sugars, e.g., sialic acids, sugar-phosphates), alditols, alcohols, and glycols are not detected or remain close to the noise level, providing selectivity of detection to the ionic and polar components of the media.  

PAD is specific for compounds, such as, carbohydrates, amino acids, alditols, glycols, and alcohols, which can be oxidized at a selected potential (applied voltage), leaving most of the other compounds undetectable, and thus, providing selectivity of detection.   

A detailed discussion on the mechanisms of ion-exchange and ion-exclusion chromatographies as well as those of SCD and PAD is beyond the scope of this article. Interested readers are encouraged to read books and publications that provide details of the principles and mechanisms of IC, SCD and PAD, including selection of columns and detectors for different types of applications (3–6). 

Fermentation broths typically have several cations and anions, including amino, carboxylic and other organic acids, as well as amines and other organic bases. However, concentrations of inorganic cations and anions are significantly higher than their organic counterparts. Thus, inorganic ions show significant peaks by IC when fermentation broths are diluted 1000-fold or more, while their organic counterparts are reduced to undetectable levels. This provides necessary selectivity for analysis.   

The first report of the determination of cations and anions using IC to follow a fermentation process of methanotrophic bacteria was published in 1992 (7). Using OmniPac PAX-500 anion-exchange column and SCD, the authors followed several anions over the course of fermentation and found acetate to accumulate when natural gas was used as the substrate, which inhibits the growth of the methanotrophic bacteria. Mono- and divalent cations were monitored by SCD using IonPac CS10 cation-exchange column eluted isocratically with 40 mM HCl–4 mM DL-2,3-diaminopropionic acid (DAP) as the eluent. The results indicated that the concentration of Mg

 + must be above 60 ppb to maintain optimum growth of the bacteria. 

Robinette et al. (8, 9) reported quantitation of eight mono- and divalent cations in fermentation broths for recombinant and pathogenic microorganisms, as well as in mammalian and insect cell cultures in complex and defined media. The authors employed IonPac CS10 and CS12 cation-exchange columns eluted isocratically with 20 mM HCl – 4 mM DAP and 20 mM methanesulfonic acid (MSA), respectively. The cations are detected by SCD. The sample preparation involves simple 1000–2500 fold dilution of filtered broth with water. The reports indicate good precision and accuracy, and a wide linear range for each cation, with detection limits ≤1 µg/mL. 

Figure 1: A time course of analysis of cations in an insect cell culture using a complex medium with glucose and sucrose as the primary carbon sources. Peaks: 1=sodium, 2=ammonium, 3=potassium, 4=magnesium, and 5=calcium.  Column: IonPac CS12; elution: 20 mM MSA, isocratic, 1.0 mL/min. (Reprinted from Ref. 9 with permission from Elsevier)  

Figure 1 shows a time course of an insect cell culture using a complex medium. The chromatograms show that the levels of sodium, potassium, magnesium, and calcium remained virtually unchanged throughout the fermentation, while ammonium concentration increased approximately 2-fold. Similar investigations were reported for 

(eukaryotic) grown in a chemically defined medium, and 

 (prokaryotic) and a mammalian cell culture in complex media containing glucose as the primary carbon source. 

Figure 2: Resolution of carbohydrates, alditols, alcohols, and glycols commonly found in fermentation broths by (A) CarboPac MA1 column and (B) CarboPac PA1 column. (Reprinted from Ref. 12 with permission from Dionex)  

Aliphatic carboxylates (e.g., acetate, lactate, pyruvate) are metabolites that often reduce fermentation yields. In an Application Note (10), Dionex reported the analysis of aliphatic carboxylates and inorganic anions in the fermentation broths of 

 in LB medium using IonPac AS11-HC column eluted with 1-60 mM NaOH and SCD. The results show well resolved peaks with good precision and accuracy, wide linear range and low detection limit for each anion. 

Figure 3: A time course of analysis of carbohydrates in H. influenzae cultivation using complex medium with glucose as the major carbon source. Peaks: 1=glucose, 2=fructose, 3=ribose; 4=maltose. H=hours.  Elution: 150 mM NaOH, isocratic, 1.0 mL/min. (Reprinted from Ref. 11 with permission from Elsevier)  

Carbohydrates are carbon sources essential for cell growth and product synthesis, while alditols and alcohols are metabolites. High Performance Anion-Exchange Chromatography (HPAEC) coupled with PAD (HPAEC–PAD) or Integrated PAD (IPAD), which is an improved PAD application, is the most widely used chromatographic technique for the analysis of carbohydrates and alditols (3–6). The CarboPac MA1 column has the ability to analyze mono- and disaccharides, alditols, and alcohols present in fermentation broths (see Figure 2A) in a single run. However, the chromatography takes as long as 60 min for each run and uses a high concentration of NaOH as the mobile phase. The CarboPac PA series of columns (e.g., PA1, PA10, PA20) take significantly less time and use low NaOH concentration. But alcohols and alditols are eluted early and are poorly resolved (Figure 2B). Therefore, the PA series columns are better suited for mono- and disaccharide analysis. Thus, the choice of column depends on the analytes to be monitored in the media. Herber and his colleagues (8, 11) used a CarboPac PA1 column to monitor microbial fermentations using chemically defined and complex media. The samples were diluted 50 fold with water and eluted from the column with NaOH (isocratic and gradient) to monitor ethanol, glycerol, galactose, glucose, mannose, fructose, raffinose, ribose, and lactose. Carboxylic acids and inorganic ions are transparent to PAD. Most of the other media components were below the detection limit at this dilution. Indeed, only proline, arginine and lysine exhibited any noticeable detector response (see later for the detection of amino acids by PAD). Figure 3 shows a time-course study of carbohydrates in a 

 fermentation in a complex medium with glucose as the major carbon source. Figure 4 shows the chromatograms of the analysis of 

fermentation broth using CarboPac MA1 (12). Glucose is the most predominant component of the media at 0 h. At the end of fermentation, no glucose was detected and the predominant components are the metabolites.  

Figure 4: Chromatograms of the analysis of S. cerevisiae fermentation broth using CarboPac MA1; (A) 0 h and (B) 24 h. (Reprinted from Ref. 12 with permission from Dionex)  

The amino acid compositions of cell culture and fermentation broths can be determined by HPAEC-PAD without sample derivatization (13–15). However, the presence of carbohydrates and alditols in the media may interfere. By optimizing the elution method, Genzel et al. (13) reported that the amino acids were well resolved and also resolved from the predominant carbohydrate, glucose, in GMEM media used in the production of inactivated influenza vaccines (see Figure 5). The authors reported good precision, accuracy, linear range, and low quantitation limit for each amino acid. Similar results were also reported for the supernatants of YPD broth, LB broth, Minimum Essential Eagle's Medium and a serum-free, protein-free hybridoma medium (14, 15).  

Figure 5: Amino acid analysis of (1) a standard amino acid mixture containing 277.5 ÂµM glucose;(2) GMEM medium; (3) GMEM medium + 1% (v/v) peptone; (4) GMEM medium+10% (v/v) FCS + 1% (v/v) peptone.  Norleucine was added as the internal standard.  Column: AminoPac PA10; elution: as per the procedure described in Ref. 16. (Reprinted from Ref. 13 with permission from Elsevier)  

Selective media are used for the growth of select microorganisms. For example, if a microorganism is resistant to a certain antibiotic, then that antibiotic can be added to the medium in order to prevent other cells, which do not possess the resistance, from growing. Selective growth media for eukaryotic cells commonly contain neomycin to select cells that have been successfully transfected with a plasmid carrying the neomycin resistance gene as a marker. Generally, neomycin is assayed following the procedure described in 

 General Chapter <81>, Antibiotics—Microbial Assay (17). However, the procedure is tedious and labor intensive, has poor precision, and requires days to complete. Neomycin is an aminoglycoside containing a carbohydrate moiety and, like other carbohydrates, can be assayed using HPAEC–PAD. Figure 6 shows the HPAEC–PAD chromatograms of two selective media containing neomycin. The media also contain glucose, sucrose, and amino acids at concentrations 100-1000 fold higher than that of neomycin. These components also show peaks in HPAEC–PAD. Thus, the resolution of neomycin was a challenge, which was accomplished using a CarboPac PA1 column and a linear elution gradient from 1.0 mM to 3.0 mM NaOH over 25 min. 

Figure 6: HPAEC-PAD chromatograms of a neomycin standard and two selective culture media containing neomycin.  Column: CarboPac PA1; flow rate: 0.5 mL/min. The effluent from the column was mixed with 100 mM NaOH before entering into the detector for baseline stabilization.  

Monitoring nutrients and metabolites in cell culture media and fermentation broth provides critical pieces of information on their roles in fermentation, and in optimizing, characterizing, and controlling the process to produce safe and efficacious therapeutic products in a reproducible manner. The results presented in this review show that the unique selectivity and sensitivity of IC with ED permit determination of fermentation broth components. This is particularly important for those components which do not have suitable chromophores for detection by absorption measurement. The sample preparation involves simple filtration and dilution of fermentation broth with water. No pre- or postcolumn derivatization is necessary for detection. In addition, the technique provides the ability to analyze the desired components selectively in the presence of others, e.g., amino acids in the presence of large excess of carbohydrates or inorganic cations and anions in the presence of amino acids. These features make IC with ED an effective technique in analysis of fermentation broths in a simple and cost effective manner.

 Parts 210 and 211, Parts 600–610 (Government Printing Office, Washington, DC). 

 2. Application Note 157, "Comparison of Suppressed to Nonsuppressed Conductivity Detection for the Determination of Common Inorganic Cations" (Dionex Corp., Sunnyvale, CA). 

Pulsed Electrochemical Detection in High Performance Liquid Chromatography (

, 4th ed., (Wiley-VCH, Weinheim, Germany, 2009). 

 6. L. Bhattacharyya and J.S. Rohrer, Eds., A

pplications of Ion Chromatography for Pharmaceutical and Biological Products

 7. L. Joergensen, A. Weimann, and H.F. Botte, 

 8. R.S.R. Robinett and W.K. Herber, J. Chromatogr. 

 9. R.S.R. Robinett, H.A. George, and W.K. Herber, 

An overview of applications in the analysis of nutrients and metabolites.

Application of Ion Chromatography with Electrochemical Detection in Optimization and Control of Fermentation and Cell Culture

Medicago has developed a proprietary plant-based manufacturing platform that produces vaccines made of influenza antigen scaffolds (i.e., viruslike particles) that are currently in clinical trials in Canada and the United States. This platform produces vaccine doses within a month of the disclosure of hemagglutinin sequences from emerging strains. It is now being automated and scaled up in North Carolina to bring its surge capacity to more than 10 million doses/month. The platform will be housed in a facility that will be built within 12 months for less than $35 million.

The recent swine H1N1 influenza pandemic revealed the limitations of the current influenza-vaccine manufacturing technologies. Although Wyman and colleagues had predicted in 2007 that egg-based manufacturing would be able to supply at least 60 million vaccines doses within five months of the declaration of a pandemic, the actual vaccine output in the 2009–2010 porcine H1N1 (pH1N1) pandemic was much lower (1). In fact, only three million doses were available five months after the identification of the causal strain. The first doses of split vaccine became available shortly thereafter, but at levels far below expectations. Fortunately, the pH1N1 strain had a low mortality rate. Had the pH1N1 pandemic had a higher severity index, the global human cost of the delays in egg-based vaccine production could have been catastrophic. Nevertheless, this outbreak has underscored the need for a faster, simpler, and more flexible manufacturing technology to respond to an emerging threat and to protect as many people as possible rapidly. 

To meet the surge capacity required for the production of pandemic vaccines, recombinant technologies are being actively developed with various expression hosts, such as bacteria, yeast, insect, or mammalian cells. Medicago developed a plant-based expression system designed to provide great surge capacity and safety, at low cost. This article will describe how this new manufacturing platform is combined with a nanoparticle antigen-presentation scaffold (i.e., viruslike particles or VLPs) that is synthesized in plants, with an extraction approach that separates the nanoparticles from the host cell components early in the process, and with a purification scheme that yields highly purified particles as drug substance.  

The authors have demonstrated that this platform efficiently produces VLPs from the sole expression of recombinant influenza hemagglutinin (HA), and that administration of these purified influenza VLPs was well tolerated and triggered an immune response of great strength and breadth in animals and humans (2, 3). The speed at which influenza VLPs can be produced following the identification of a new HA gene sequence was assessed in the context of the pH1N1 pandemic. The first doses of a plant-made H1 VLP candidate vaccine against the pH1N1 strain were produced within three weeks of the disclosure of the new HA sequence by the Centers for Disease Control and Prevention (see Figure 1).  

Figure 1: H1 viruslike particle (VLP) experimental vaccine production timeline. (FIGURES ARE COURTESY OF THE AUTHORS)			

The manufacturing platform based on transient expression, and the VLP-based antigen-presentation platform were developed in the context of the pandemic threat. They now have been applied successfully to the development of a trivalent seasonal vaccine. Vaccine products for these two indications have been produced under good manufacturing practice (GMP) conditions and currently are in clinical trials in Canada and the United States.  

The transient expression system uses the capacity of a bacterium, 

 to infect plant cells and transfer essential genetic information in the form of a mobile DNA fragment to the nucleus of the infected cell. Because this mobile DNA fragment only remains viable for a few days in the nucleus and is not integrated in the plant genome, the cells are only transiently transformed. Despite the transient nature of the transformation, this technology results in extremely high expression of the gene of interest for days after cell infection, and enables synthesis of large quantities of the targeted protein. 

Medicago's transient expression system starts with the assembly of an expression cassette where the target gene is positioned within transcription and translation components adapted for expression in plants. The expression cassette is inserted within mobile DNA borders of a plasmid, which is transferred into the bacterial transfection vector 

 Transfection of the mobile DNA copy containing the expression cassette to 

 leaf cells is performed by soaking healthy plants upside down in a vacuum tank containing a liquid suspension of 

 the inoculum. As vacuum is applied, the air-filled cavities inside the leaves are emptied, thus leaving space for infiltration of the bacterial inoculum upon release of the vacuum. As the inoculum is infiltrated, the 

 establish contact with parenchyma cells, which constitute the vast majority of the leaf active cells. Because vacuum infiltration is instantaneous and highly invasive, infection of leaf cells is also instantaneous, which helps to synchronize cell infection and the resulting transfer of mobile DNA copies containing the expression cassette. Transient expression is extremely fast and results in high accumulation of the recombinant protein within three to six days. 

In the life cycle of the influenza virus, formation of the viral particle involves the budding of a portion of the host plasma membrane saturated with the viral surface proteins. HA is the major surface protein of the influenza virus with a transmembrane domain that spans through the viral envelope. The authors demonstrated that transient expression of the HA surface protein in plant cells results in the formation of VLPs budding from microdomains of the plasma membrane saturated with recombinant HA.  

At maturity, these VLPs have morphological features similar to those of the influenza virus, but only contain its immunogenic determinants, with none of the viral RNA, which is packaged within the enveloped particles of true viruses. Compared with isolated soluble antigens, antigens at the surface of VLPs have the double advantage of adopting a conformation that is identical to those found on the cognate virus, and of being presented in a multivalent structure similar to that of the native virus. These features enable VLPs to achieve enhanced stimulation of the immune response with balanced humoral and cellular components. This benefit of VLPs over soluble recombinant antigens is believed to be particularly strong in vaccine products against enveloped viruses because VLPs present the surface antigens in their natural, membrane-bound state (4). As the influenza viral particle, influenza VLPs are enveloped, pleiomorphic, and 100 to 150 nm in size (see Figure 2).  

Figure 2: Structural characteristics of influenza virus and viruslike particles, including (a) a comparison of surface attributes, (b) a cross-section showing internal distinctions, and (c) transmission electron microscopy images of influenza viral particles and influenza viruslike particles.			

HA expression plasmid, master, and working cell bank of transformed Agrobacteria

The recombinant influenza HA gene is built in such a way that native HA is expressed, thus yielding a mature protein product identical to the wild type influenza HA protein. Expression is maximized by the integration of elements that result in high transcription of mRNA and hypertranslation of the product. In addition to these elements, a suppressor of silencing is coexpressed with the HA protein to counteract specific HA mRNA degradation (i.e., mRNA silencing) that is triggered by the significant accumulation of HA mRNA in plant cells.  

 the plasmids are fully sequenced to ensure sequence integrity. Upon confirmation of integrity, plasmids are introduced into 

 selected clones are grown in a master cell bank (MCB) that generates the working cell bank (WCB). The MCB and WCB are tested for purity and plasmid integrity. 

A liquid culture of the bacterial vector is produced from the WCB and used to prepare the 

 is a wild Australian relative of cultivated tobacco (i.e., 

 plants are grown in controlled and contained conditions. A high-quality, soilless, sterilized root-substrate medium provides physical support and optimal water retention. Apart from fertilization and substrate moisture, which are adjusted during plant development, all growth conditions remain unchanged during the production cycle (i.e., 40 days). All components used for biomass production are released under standard operating procedures. Special care is taken to control the presence of heavy metals in all liquids and solids that contact the biomass. Fungal, microbial, and algae growth are strictly controlled during all phases of growth. 

Plant development is monitored from seeding to infiltration to ensure proper and reproducible biomass development. Plants are characterized during their growth and must meet specific criteria to be accepted for inoculum infiltration. 

The authors' pilot facility in Québec City, Canada, has an inoculum infiltration capacity of 1200 plants per day. Infiltration is performed in a stainless-steel tank, 15 plants at a time. The vacuum infiltration lasts only 1 min, and the procedure is computer-controlled to ensure reproducibility. At a plant under construction in North Carolina, the infiltration unit will accommodate 15,000 plants per day. Transportation of the biomass to and from the infiltration unit will be completely automated. Following infiltration, plants are again maintained in controlled and contained conditions during the three to six days required for transient expression to take place. 

Plant leaves are collected and mixed with a depolymerization solution. Medicago has developed a proprietary technology that treats the biomass with cell-wall depolymerization agents. This treatment acts on structural or cross-linking components of the cell wall, which loosens the rigid extracellular matrix of the leaf cells, and subsequently releases the VLPs into the solution. The procedure releases more than 90% of intact VLPs with less than ~5% of the host-cell components (i.e., protein and DNA), and is thus an extremely efficient primary recovery step. 

Because VLPs are large particles (130 nm in diameter), they can be concentrated by tangential-flow filtration (TFF). This concentration step also eliminates most of the remaining host-cell components, including low molecular-weight compounds. Diafiltration at this stage conditions the VLPs in preparation for chromatography. 

VLPs are purified by ion-exchange chromatography, concentrated, and diafiltered against the formulation buffer through a final TFF. Chromatography also allows the removal of residual DNA and endotoxins. 

CHARACTERISTICS OF PURIFIED INFLUENZA VLPS

The purified influenza VLP vaccines used in clinical trials were released under the criteria used for licensed influenza vaccines and have met all standards for purity and identity. DNA and endotoxins were always below acceptable limits, and these preparations were tested for sterility and subjected to General Safety Testing in guinea pigs before being released. 

Table I: Characteristics of purified H5 viruslike particles (VLPs).			

Mass spectrometry helps to identify trace protein contaminants. Most of the contaminants are naturally associated with plant plasma membrane and are neither immunogenic nor induce an allergic response in animal and human models. Because this product arises from a new platform, an extensive complementary physicochemical characterization was performed on purified preparations. Table I presents the measured attributes of the pandemic H5 VLP vaccine product analyzed on multiple lots. The narrow range of measurements for mean particle size, HA purity, and lipid–protein ratio demonstrates the high lot-to-lot reproducibility of the manufacturing process. The Coomasie-stained SDS-PAGE and Western blot analysis of the pandemic H5 VLP vaccine product in Figure 3 further illustrate the purity level of the product (i.e., 98–99% purity). 

Figure 3: Purity and identity of an H5 viruslike-particle (VLP) vaccine: Coomassie-stained SDS-PAGE of (1) molecular-weight marker and (2) 2.5-Î¼g of influenza H5 VLP vaccine, and Western blot analysis using polyclonal antibodies against influenza H5N1 virus (strain A/Indonesia/5/05) of (3) 50 ng and (4) 500 ng of the influenza H5 VLP vaccine.			

The pandemic H5 VLP vaccine product has been tested for safety and immunogenicity in a completed Phase I clinical trial and an ongoing Phase II trial in Canada. The Phase I results demonstrated that the vaccine product was safe and well tolerated and did not induce allergenic responses. No increase in the level of naturally occurring serum antibodies to plant N-glycans was observed. The evaluation of immune response to VLP immunization by hemagglutination-inhibition, single-radial hemolysis, and microneutralization assays were well correlated and showed a clear dose-response (3). In the Phase II trial, the preliminary results showed 65% seroprotection and 65% seroconversion after the administration of two 20-μg doses to 18 to 49 year-old subjects.  

Today, the globalization of trade and the amount of international travel provide many opportunities for infectious diseases to spread rapidly and globally. As seen with the outbreak of the 2009 A–H1N1 influenza, within two days of the first confirmed cases from Mexico, the virus was reported in five additional countries. Because of difficulties in producing the vaccine, initial doses became available only after 26 weeks in the US, and supplies for the protection of all US citizens would have taken almost a year to produce. The President's Council of Advisors on Science and Technology stated that more than 2000 lives—with an average age of less than 40—could have been saved if vaccination had begun even one month earlier (5). 

This increasing risk is posing new challenges to public-health agencies around the globe and stressing the need for effective vaccines sourced from highly flexible manufacturing facilities that can be mobilized rapidly and cost-effectively. The greatest potential to meet this challenge lies in the use of recombinant DNA technologies.  

Medicago is pursuing a promising alternative approach that combines the speed of transient-expression technology with the efficacy of VLPs as antigen-presentation scaffolds. The company's manufacturing platform offers surge capacity and cost advantages over recombinant technologies, such as mammalian and insect cells. The technology can deliver a vaccine for testing in less than a month after the disclosure of genetic sequences from a pandemic strain, as demonstrated by the first doses of plant-made H1 VLP candidate vaccine that were produced at the onset of the 2009 pandemic. This production time frame thus has the potential to allow the vaccination of a population before the first wave of a pandemic strikes, and therefore to supply large volumes of vaccine to the world market. 

In the past five years, Medicago has successfully developed this technology from the bench scale to the pilot scale according to cGMP. The compnay is now transitioning into large-scale manufacturing. Since 2008, Medicago has operated a 24,000-ft

  pilot clinical cGMP facility located in Québec City. This facility includes 10,000 ft

  of high-tech contained plant biomass production space, as well as 14,000 ft

  of cGMP manufacturing units for plant manipulation, product recovery, and purification. 

In August 2010, Medicago began building a 97,000-ft

  cGMP facility in Research Triangle Park, North Carolina. Designed to produce 10 million doses of pandemic influenza vaccine per month, this facility will include a fully automated and contained plant biomass production facility and a state-of-the-art extraction-and-purification unit. On an annual basis, this facility would have a production capacity of 40 million doses of seasonal-trivalent influenza vaccine or 120 million doses of pandemic-influenza vaccine. The facility will be built in a 12-month timeframe at a cost of less than $35 million.  

The capacity, speed of construction, and low cost of Medicago's large-scale facility clearly illustrates the promises of the company's manufacturing platform. These factors, in addition to the interim Phase II clinical results of Medicago's first enveloped VLP vaccine, could form the basis for a fast, high-quality, and cost-effective manufacturing solution to address the needs for improved 21st-century vaccines.  

 is vice-president and chief scientific officer, 

 is vice-president of product development, 

 is vice-president of business development, 

 is president and chief executive officer, all at Medicago, 1020 Route de l'Eglise, Suite 600, Québec, QC, Canada, G1V 3V9, 

1. O. Wyman and the Program for Appropriate Technology in Health, "Influenza Vaccine Strategies for Broad Global Access," (PATH, Washington, DC, 2007). 

A plant-based expression system could provide greater speed and capacity at a comparatively low cost.

Plants As an Innovative and Accelerated Vaccine-Manufacturing Solution

Hollow fiber bioreactors have been in use for more than 30 years for the production of secreted proteins and antibodies from mammalian cells. Advances in fiber construction and methods for oxygenation have the potential to bring the advantages of hollow fiber cell culture to biomanufacturing. 

 cell growth conditions, Richard Knazek developed the hollow fiber bioreactor (HFBR) in 1972 (1). The HFBR system is a high-density, continuous-perfusion culture system.  It is a well-established system for the production of secreted proteins from hybridoma, Chinese hamster ovary cells (CHO), human embryonic kidney (HEK) 293 and other mammalian and insect cells. The primary advantage is that it is possible to culture large numbers of cells at high density, resulting in product accumulating to very high concentrations. Up until now, the application of HFBR technology has been limited to laboratory scale applications. HFBRs provide some features that are advantageous in biopharmaceutical manufacturing, such as continuous, modular, flexible manufacturing formats employing compact and disposable systems. There is also growing interest in perfusion-based bioproduction protocols (2). FiberCell Systems has developed a large-scale HFBR system that brings the benefits of hollow fiber cell culture to the bio-production arena (see Figure 1). 

Figure 1: Cells growing at high density in Fibercell Systems hollow-fiber bioreactor (HFBR). 			

HFBRs consist of thousands of semi-permeable hollow fibers in a parallel array within a tubular housing or cartridge, fitted with inlet and outlet ports. These fiber bundles are potted at each end so that any liquid entering the ends of the cartridge will necessarily flow through the interior of the fibers. Cells are generally seeded within the cartridge, but outside of the hollow fibers in what is referred to as the extra capillary space (ECS) (see Figure 2). Culture media is pumped inside the hollow fibers, allowing dissolved gas, nutrients, and waste products to diffuse across the fiber walls. Once having passed through the cartridge, the culture medium is oxygenated and returned to the cartridge.  

Figure 2: Schematic of the growing conditions within a HFBR. 			

Unlike current popular batch-style manufacturing formats, including such disposable systems as rocked bags and bags supported by stainless steel containers, HFBRs do not subject cells to significant variations in primary and secondary metabolites (such as glucose and lactate), oxygen, or product levels. A HFBR significantly cuts the operating footprint and reduces the product harvest volume by factor of 10 to 100 times. Furthermore, it avoids such scale-determined discrepancies as impeller and sparge-induced hydrodynamic forces, and reactor column height-induced differential gas solubility. The high cell density and low-sheer environment permits the use of simplified serum-free, protein-free media that can be less expensive and that does not require such components as synthetic membrane protective surfactants or antifoams. Attached cultures are supported, therefore no lengthy adaptation of cells to suspension growth is required.  The 10–100 times more concentrated secreted product, the absence of high molecular weight medium components, and the greatly reduced levels of host-cell proteins and DNA facilitate downstream processing, a major cost in bioproduction. 

Bringing the advantages of HFBRs to large-scale bio-manufacturing has long been a goal for those familiar with hollow fiber systems. The operational scale for HFBR systems has been limited by several factors: 					

Solubility of gas in cell culture medium decreases as the temperature is increased to 37° C. 

It is difficult to design a pump that will provide 6–10 L/min flow rate without denaturing biological components or generating significant backpressure.

The gross filtration rate of cellulosic hollow fibers (i.e., the rate of nutrient and waste product exchange) have typically been too low to effectively support the culture of adherent cell lines such as CHO and HEK293.

Fiber geometry is such that the fibers were not uniformly distributed within the cartridge housing, resulting in areas where a large inter-fiber distance led to necrotic areas within the cartridge.

FiberCell Systems has developed a unique approach to address these issues. Instead of using pumps and systems to perform gas exchange with the cell culture medium, a gravity-based system for medium flow that also directly oxygenates the bioreactor cartridge from the inside out has been developed (see Figure 3). A new generation of high flux fibers is employed, and they are also cast with "waves" in them to evenly distribute the fiber bundle within the hollow fiber cartridge. With a cartridge volume of 1 L, this fully disposable system has the potential to replace up to 10,000 L stirred tank reactors for mammalian protein production over a 100–day period of culture. This large-scale system offers all of the advantages outlined above, along with others that are specific to bio-production in a disposable system. 

Figure 3: Fibercell Systems prototype large-scale HFBR. 			

There are four fundamental characteristics that differentiate hollow fiber cell culture from classic suspension culture: 					

Cells are bound to a porous matrix much as they are 

Cells are cultured in a gently perfused, non-shear environment.

The molecular weight cut off of the support matrix can be controlled.

The system possesses an extremely high surface area to volume ratio (150 cm

Cells are bound to a porous support much as they are

There is no requirement to split cells. Cells in this perfusion system maintain viability and production-relevant metabolism in a post-confluent manner for extended periods of time, months or longer. Passage number is basically irrelevant and a certain percentage of the cells will become quiescent, that is, they will produce protein but do not divide.  For example, one hybridoma was reported to maintain efficacious productivity for over one year of culture while a glioma cell line was cultured for nearly two years of continuous HFBR culture. 

Cells are cultured in a gently perfused, non-shear environment.  

Unlike suspension culture, the cells are constrained within a non-shear compartment with gas, nutrition and waste transport moving across the fibers. Lack of shear simplifies the medium composition, and perfusion permits a more homeostatic, 

-like environment for the cells. This can result in improved protein translational fidelity. These growth conditions and lack of shear also result in significantly reduced apoptosis (3). The majority of cells that become necrotic will not release host cell proteins or DNA into the culture medium, resulting in a product that is cleaner and easier to purify from the bulk harvest 

The molecular weight cut off of the fiber can be controlled.

Desired products can be retained to 10–100 times higher concentrations. Toxic or unstable products can be selectively removed, and the effects of cytokines controlled.  An example of this is hybridoma culture, in which the inhibitory cytokine transforming growth factor beta can be selectively removed from the culture, while the secreted antibody is retained.  

The system possesses an extremely high surface area to volume ratio.  

HFBRs have an extremely high surface area to volume ratio (in the range of 100–200 cm

 /mL of volume). Coupled with the high gross filtration rate of FiberCell Systems' polysulfone fibers, the exchange of nutrients and waste products is rapid. Cell densities of 1–2 × 10

 tissue-like densities. A module with a volume of 1 L would support as many cells as a 100 L stirred tank, as well as providing continuous production. High cell densities support higher volumetric productivities, and facilitate adaptation to lower serum concentrations or serum free culture. In fact, a simplified, protein-free, chemically defined, animal component-free serum replacement, CDM–HD has been developed by FiberCell Systems and is optimized for use in hollow fiber systems.  The use of a protein-free medium such as CDM–HD provides cleaner harvests of product, easier regulatory compliance and simplified downstream purification.  

-like cell culture conditions can also result in improved protein folding and more uniform glycosylation patterns over time. Since it is a continuous perfusion system, the amount of protein produced is determined as much by the length of time the culture is maintained as by such parameters as the clone's specific productivity or the size of the cartridge.  

The combination of an unlimited nutrient supply and the ability to de-bulk the culture through the cartridge ports allows the system to be maintained at relative equilibrium for several months or longer. This continuous production over long periods of time, rather than the severely batch-style results from other systems, provides several benefits, including: consistency in culture condition, dramatically increased production per unit footprint and culture volume, continuous or daily product harvest allowing timely and convenient stabilizing treatment or storage conditions, and products that might be toxic or inhibitory to cells can be selectively removed from the culture. 

The system included the following components: 					

A disposable, preassembled, and sterilized package including the cartridge, two media bags with fittings, and two disposable sensor "windows" with reactive pads bound inside.

Gravity pump with motorized cartridge angle controller.

The diameter of the fibers has been increased to reduce the resistance to flow while still presenting a large surface area to volume ratio.  This allows the use of a gravity pumping mechanism that can generate a flow of 5–10 L of media per minute through the interior of the hollow fiber bundle. The gravity pump will then simply reverse direction. The bags contain both medium and gas, and when the liquid is finished flowing through the cartridge, the gas is drawn through the interior of the fibers, directly oxygenating the cartridge from within.  The disposable bags (Thermo Scientific HyClone BPCs employing CX5-14, a pentalaminate animal component-free film) are gas-impermeable, and the gas composition inside the bags is tightly controlled for optimum cell culture performance. The environmental enclosure is a simple, non-pressurized, temperature controlled incubator, but can be modified to include further environmental control such as HEPA filters to create a Class 100 clean room environment within the system for cGMP operation. Sensors for pH, O

, and potentially real-time glucose monitor the culture environment. The absence of cells in the medium bags to be monitored supports more robust and sensitive measurements. These sensors will support the control of medium and gas in the media bags. More advanced monitoring is possible using any one of a number of new approaches, including capacitance based monitoring probes.  This exciting new technology employs passive electrical (dielectric) radio frequency-based sensors to measure the overall capacitance, and therefore viability, of the cell mass within the cartridge. The low volume in both the cartridge and the medium present in the system at any one time makes both monitoring and process control more precise and responsive, even though the system may consume 50–100 L/day.  Stacking like-sized cartridges with their media bags on top of each other can provide additional scale. 

Figure 4: Hollow fiber cartridge on Fibercell Systems prototype large-scale HFBR. 			

The FiberCell Systems HFBR allows the production of about 20–200 mg of antibody per week using the 20 mL cartridge (FiberCell Systems catalog # C5011). The product is harvested in a concentrated form 500–5000 µg/mL in a volume of 20–40 mL and production of the antibody can be accomplished in a standard CO

As an example, two monoclonal antibodies produced in DMEM + CDM-HD are shown in Figure 5. Length of culture was three weeks, total antibody produced was in excess of 140 mg each, and average concentration was 3.0 mg/mL. The dialyzed supernatants had their protein concentration quantified and purity check on a 12% sodium dodecyl sulfate polyacrylamide gel. A total of 12 L of medium per antibody was consumed. 

Figure 5: Raw supernatant harvest was briefly dialyzed to reduce sample tonicity. No other purification was performed. The heavily loaded samples demonstrate low levels of contaminating proteins. (Data courtesy of Dr. Erin Bromage, U. Mass., Amherst) 			

Extrapolating this data to a 1 L sized hollow fiber bioreactor cartridge would result in total antibody production of approximately 3 g/day in a volume of 1 L, consuming 50 L of DMEM/CDM-HD per day. 

473 mg of purified recombinant protein recovered from an rCHO cell line was harvested from the FiberCell Systems 20kd MWCO cartridge (FiberCell Systems catalog # C2018). Medium was Dulbecco's Modified Eagles Medium (DMEM) with 2% fetal bovine serum, each harvest was 70 mL in volume, and total harvest volume was 4.8 L, for an average protein concentration of approximately 100 µg/mL/d. The protein was a very complex, hexamerized immunoglobulin G (IgG) consisting of 6 IgG1 subunits held together with three IgA tails. The variable fragment (Fv) region was modified to contain the CD4 receptor (2). The cartridge consumed an average of 2 L of medium per day over a 60–day period of production. 

An interesting observation was the comparison of protein produced using T–flasks versus the hollow fiber cartridge (see Figure 6). When produced in flasks, approximately 40 % of the protein was secreted as an unfolded monomeric subunit. Placing the same cells into the HFBR cartridge resulted in nearly 95 % of the protein being produced as a properly folded hexamer (3). Better cell culture conditions resulted in better protein expression fidelity. 

Figure 6: Gel filtration chromatography of a hexeramized recombinant immunoglobulin (IgG). When cultured in flasks (top trace), approximately 40% of the protein is expressed as an improperly folded monomeric subunit (B) rather than the hexamer (A). These cells when transferred to a hollow fiber bioreactor exhibit 95% complete folding ( C ) as a result of the improved cell-culture conditions.			

Expression levels were significantly below those found in commercially optimized CHO cell lines, although the results are still impressive. A 1 L sized HFBR cartridge would have a  total harvest volume of 100 L, total protein recovered would be 10 g and total medium consumed would be 2,400 L. 

246 mg of purified recombinant IgG1 from a CHO (DG44) cell line was harvested from the FiberCell Systems 20kd MWCO cartridge (FiberCell Systems catalog #C2011). Medium was a serum-free, protein-free formulation similar to CDM-HD. Each harvest was 20 mL in volume; total harvest volume was 320 mL for an average concentration of over 800 µg/day/mL. The cartridge consumed 1 L of medium every three days and the culture was maintained for a total of 35 days.  For technical reasons, the cell viability in this run dropped rather low, however, the harvested protein produce was remarkably uncontaminated, as demonstrated by the gel of the unpurified harvest in Figure 7. In a  1 L  sized HFBR, the total protein recovered would be 12.5 g in a volume of 16 L with 500 L of medium consumed.  

Figure 7: Harvest from bioreactor rCHO cells. Raw supernatant was briefly dialyzed to reduce sample tonicity, no further purification was performed. 			

To extrapolate these results to a commercially prepared recombinant CHO cell line with expression in the range of 1 g/L in a standard stirred reactor, the 1 L sized HFBR cartridge should be capable of producing 10 g or more of protein per day, while consuming 100 L of medium. 

A hollow fiber matrix allows for efficient harvest of secreted proteins.

The Potential Application of Hollow Fiber Bioreactors to Large-Scale Production

Issue PDF

Advances in Fermentation, Cell Culture, and Expression Systems

BioPharm International-05-02-2011

Increasing Efficiency in Protein and Cell Production by Combining Single-Use Bioreactor Technology and Perfusion

Application of Ion Chromatography with Electrochemical Detection in Optimization and Control of Fermentation and Cell Culture

Plants As an Innovative and Accelerated Vaccine-Manufacturing Solution

The Potential Application of Hollow Fiber Bioreactors to Large-Scale Production

BioPharm International, May 2011 Supplement (PDF)