Throughout the ages, great sculptors like Michelangelo have said that their work is about taking a block of marble and freeing a form that is already there. All the artist needs to do is remove everything that does not belong in the masterpiece. 

The work of downstream processing scientists is similar. Instead of stone, their artistic medium is a colored, and sometimes turbid, soluion. Purification must remove process contaminants and produce a pure end product. It is that simple. 

In reality, there is a conflict between upstream and downstream processes resulting from the industry's strength in fermentation and weakness in purification. The consequences are dramatic. Monoclonal antibodies (MAbs)—the fastest growing biopharmaceutical market segment, with predicted annual sales of $20 billion by 2010—are an example. Demand has already reached 6 metric tons this year. 

Fortunately, large amounts of raw material are accessible as a result of sufficient manufacturing capacity that has become available recently, combined with significant productivity increases in upstream processes.

  Experts claim that even after recent dramatic improvements from mg/L to g/L of antibody titers in mammalian cell culture, the best is yet to come.

  From the early days of MAb manufacturing, we know that physiological levels of up to 20 g/L can be achieved in mouse ascites. That is probably the long-term goal we are looking at. 

Unfortunately, we have not seen the same speed of progress further downstream. Downstream operations have always been in a sandwich position. Typically, senior management focuses on the initial phase of the process, where most investment goes and where the raw material is generated. The other typical focus is formulation, as the outlet for the drug product. For years, downstream simply had to deliver; nobody asked how. Now, as the bottleneck becomes obvious, there is a lot of pressure on purification. We are in a technology crisis, and protein purification must make a comeback.

Today's downstream processing operations generally focus on two main areas: a) the initial recovery phase, including capturing, where bulk purity is achieved; and b) the subsequent polishing phase, which adds safety through orthogonal strategies for impurity and pathogen clearance.  

It is relatively easy to identify the bottlenecks in the process. Early recovery, including harvesting, cell removal, and clarification, is not a problem. These steps can be operated with robust technology that processes high volumes in the biopharmaceutical industry; and even larger volumes—beyond the 100,000-L scale—in food, beverage, and technical enzyme manufacturing. 

But further downstream, for example in capturing, processes are dominated by chromatography with its inherent limitations. Very large scale chromatography is technically feasible, but big columns encounter big problems and current upstream titer developments are pushing biochromatography beyond its physical and economic limits. 

Protein A is the work horse in antibody separation, and accounts for up to 10% of the overall costs; individual column fillings cost millions of US dollars but still cannot cope with the volume of common fermenter batches. Despite such problems, however, Protein A offers unmet specificity, so it will continue to be the industry standard until a breakthrough arises from the second or third generations of the mimetic ligands that are currently available. 

Because state-of-the-art capturing, including affinity chromatography, yields material that is >98% pure, polishing efforts focus on removing pathogens and trace contaminants. Because this area does not receive much attention, polishing does not seem to represent a bottleneck. In reality, it does, because in the standard approach— removing typical process-derived contaminants with anion exchangers in flow-through mode—the flow rate, rather than binding capacity, becomes the limiting factor. Thus, we run into a problem: to increase the flow rate, we need very large chromatography columns, which require very large quantities of expensive resins. 

Fortunately, the polishing area—unlike the capture area—is beginning to benefit from advances such as disposable chromatography systems, which have been implemented in a number of processes.

  In one example, a disposable 0.5-L membrane can remove process-derived impurities such as DNA, viruses, and endotoxins from a 1000-L feed stream just as well as a stainless steel column that is a hundred times larger. Results from a case study examining the economics of such systems demonstrated that disposable equipment can provide a cost savings up to 70% in MAb polishing.

  The disposable options benefit from lower capital investment and significantly reduced material costs because these small devices consume 95% less buffer. And for viral clearance, disposable chromatography offers additional advantages beyond cost savings: it eliminates the need to carry out virus carryover and cleaning validation studies. 

In many processes, virus filtration (the other main method for virus removal) is the most expensive downstream step, accounting for up to 40% of costs. Thus, it is understandable that this step is a target for process optimization. Overall productivity, however, is not only a matter of initial flow rate, but also of total throughput and sensitivity to aggregate and impurity level. A viral clearance study with at least four viruses for late-stage development is state-of-the-art, and log reduction values (LRVs) of 4 and higher are hygienic factors. More interesting, but also less trivial, are LRVs that are independent of the flow decay of the filter. 

A clear trend in bioseparations is to combine steps, even eliminating the need for staging tanks and hold-up volumes between unit operations. Orthogonal strategies for each objective are key to achieving industry goals for yield and quality.  

With increasing biomass in cell culture, high-productivity harvesting procedures are enticing, and there is a lot of interaction among the three unit operations of centrifugation, cross flow, and depth filtration. Eventually, we will see more integrated processes, such as expanded bed adsorption, ideally in disposable formats.

 For monoclonal antibody manufacturing, for example, we need to develop processes that require only two column steps followed by a rational filtration train, in which every step addresses the removal of a certain contaminant. 

Polishing before capturing sounds provocative, but is becoming more common. This approach protects the protein A column and prevents the column eluate from precipitating by removing certain lipoproteins from the feed stream. These concepts can be custom designed with smart membranes that remove individual contaminants like proteases very specifically and early in the process to yield stable storage forms.  

In chromatography, attempts are being made to make resins look more like membranes and vice versa. It makes much more sense to accept the limitations of both technologies. They each are great in their individual core application areas and deserve an integrated approach, for example, by using a chromatography train (capturing phase) followed by a filtration train (polishing phase). Capturing will always require high dynamic capacity and will be dominated by resins (with the exceptions of large molecules and high dilutions), whereas polishing is going to be the domain of charged membranes. 

Thus, we are seeing many paradigm shifts in bioseparations, including the trend toward disposables to save costs. I predict that in a number of years, polishing applications will be dominated by disposable concepts. But much more is required. If downstream processing had kept pace  with the revolution in fermentation, chromatographic separations would require only minutes, not hours. Because this is not feasible, we must seek alternatives. Bioseparation is desperate for high-throughput, high-productivity unit operations. One place to start would be to copy what has been done in food, beverage, and technical enzyme manufacturing.  

In sum, robust and reliable methods such as extraction, precipitation, and crystallization deserve a renaissance. Michelangelo, although considered a master, was said to be constantly dissatisfied with himself. He probably realized that complacency is a dangerous enemy. We should take our cue from him. 

1. BioPlan Associates. 3rd annual report & survey on biopharmaceutical manufacturing capacity and production. June 2005. 

2. Wurm FM, Production of recombinant protein therapeutics in cultivated mammalian cells. Nature Biotechnol. 2004; 22: 1393–1398. 

3. Aldridge S. Downstream processing needs a boost. GEN 2006; 26(1). 

4. Zhou JX, Tressel T. Basic concepts in Q membrane chroma-tography for large scale antibody production. Biotechnol. Progress. 2006; 22(2):341–349. 

5. Sinclair A. Disposable technologies—impact on biomanu-facturing. London: Institution of Chemical Engineers; 2005. 

Articles

The polishing process-unlike capturing-is beginning to benefit from technological advances.

The Renaissance of Protein Purification

Traditional chemical analysis in downstream biotechnology manufacturing operations has been performed in off-line laboratories which are physically and organizationally remote, from the manufacturing site. Results of these assays are often not provided in a timeframe that is useful in controlling and monitoring the downstream process with maximum efficiency. Reports of delays (ranging from hours to days) in obtaining results of analytical monitoring of downstream processes often relegates these functions and the corresponding data to a compilation of historical databases rather than to its intended use as a tool to manage and control the real-time process under observation. Modern manufacturing processes in other industries, such as semiconductor and petrochemical processing, have long recognized and employed the advantages of performing analytical measurements at the production line and have widely used these techniques to enjoy advantages in production efficiency and product quality. The comparison and contrast to the current state of biotechnology manufacturing process efficiency and quality is particularly evident. FDA has long considered application of on-line analytical techniques as credible opportunities to improve product quality and efficiency in pharmaceutical production, including biotechnology and biological production. As a result, FDA began investigating technologies and prospects for application of automation in pharmaceutical, biotechnology, and biological manufacturing processes in ten years ago. 

The purpose of the PAT initiative is to move analytical laboratory functions close to the manufacturing process and to improve manufacturing efficiencies and product quality. This would be accomplished by providing real time support and control of manufacturing processes through analysis of the process stream coupled with statistical process control and tight feedback control loops. 

The FDA PAT initiative was created formally by the Center for Drug Evaluation and Research (CDER) branch of the FDA in 2002

  and sought to provide a framework to employ these techniques for small molecule pharmaceutical production under guidance from FDA. PAT processes

  are becoming an established tool to promote and improve quality and production efficiency in the pharmaceutical manufacturing process. The successes of early implementation in small molecule manufacturing processes led early adopters in the biotech community to explore PAT techniques and processes for potential use in biotechnology applications. It has been demonstrated that the same PAT technology used in traditional pharmaceutical manufacturing can be employed in biotechnology manufacturing with the same positive effect. To that end, FDA is now focusing its PAT initiative on biotechnology and biologicals manufacturing through its Center for Biologics Evaluation and Research (CBER) and PAT offices. 

Successful PAT programs are defined by a careful combined application of statistical process control (chemometrics) and use of on-line, at-line, or near-line sensors and instrumentation capable of measuring key parameters of the manufacturing system in real time. These capabilities are arguably most critical and useful in the final stages of biotechnology manufacturing, where the value of the product is at its highest. Downstream processing applications are a specific target for implementation of PAT processes because this production stage is where the desired product of the fermentation or cell culture process is separated from the complex production matrix, then concentrated, identified, quantitated, and collected for final processing and formulation. These operations are particularly suited for use of on-line, at-line, and near-line sensors and analytical instrumentation associated with corresponding chemometric and process control systems. 

Statistical process control and chemometric systems are employed across a broad spectrum of industries and are not further explored here. References to general chemometrics and chemometrics specific to biotechnology and biologicals are widely available.

  The focus of this article is on-line sensors and instrumentation specific to downstream processing. 

The primary analysis requirement in biotechnology downstream processing is molecular identification and quantitation; used to identify and quantitate the product to determine its purity after separation from its production matrix by process liquid chromatography. Therefore, the most powerful and useful analytical techniques to consider for downstream processing in the PAT context are spectroscopy and analytical scale separation methods that can characterize analytes on a molecular basis. A typical downstream process scale chromatographic technique not only purifies but concentrates the analyte and is operated at high load factors and high flow rates, therefore, the associated analytical technique must have a wide dynamic range and a fast cycle time. Typical cycle time requirements for on-line downstream assays are under 15 min, with many under 10 min. This cycle time requirement plus the need to separate, characterize, and identify the desired product in an often complex matrix composed of chemical species similar to the desired product,  requires precise, high speed, purpose-designed instrumentation characterized by high accuracy, reproducibility, and resolving power. This combination of requirements limits the discussion of applicable downstream biotech PAT instrumentation to near infrared spectroscopy (NIR) and other specialty optical spectroscopy techniques, such as mass spectroscopy, liquid chromatography, and electrophoretic techniques. 

NIR has enjoyed some success in pharmaceutical manufacturing operations, primarily in blending and formulations of dry bulk materials. Its use in on-line downstream processing applications for identification and quantitation of product is less established, but its possibilities should not be overlooked. Practical on-line NIR analyses for biotechnology applications have been demonstrated.

  Factors that will control use of NIR for downstream process applications include probe design and placement, calibration methodology, and heavy reliance of NIR on statistical analysis to calibrate and report concentration and identity of analytes. In many applications, a major advantage of NIR is non-contact. In these applications, the spectrometer is not in physical contact with the sample and, therefore enjoys long and reliable operation. In the case of a downstream process application however, the spectrometer samples the analyte by inserting an optical probe into a product transfer line at an appropriate sample point. This has the advantage that the sterility of the downstream system is maintained and that there is no physical withdrawal of sample from the process line. However, with the probe in direct contact with the fluid sample matrix, proper orientation and maintenance of this probe is critical to the technique's success. Fouling resulting from deposition of biological material on the probe quickly renders the measurement unusable. Furthermore, the low frequency characteristics of the spectra produced in NIR requires heavy reliance on calibration by multivariate solution of simultaneous equations. The sample and its matrix must be fully characterized and non-variate for the calibration to remain valid. This is not a simple task in a complex sample and matrix characteristic of biotechnology production. The speed and simplicity of instrumentation do, however, make it appropriate for consideration for downstream process applications and early applications of this technology appear promising. 

Mass spectroscopy (MS) is a powerful and well appreciated tool, finding broad acceptance in a number of non-traditional applications including biotechnology. For example, mass spectrometers have been installed and used in upstream applications for measurement of headspace gas analysis in cell culture and fermentation reactors, to monitor O

. Mass spectrometry can be used to determine absolute analyte identity and for quantitation—both desirable attributes. It is rapid and has high resolving capability. In downstream processing, a mass spectrometer could however,  acquire a sample stream from an automated sample port or sampling system at the appropriate point in the process stream. Mass spectrometer applications in a downstream process are effectively prohibited by typical high salt content found in many biotech process streams. All known commercial fluid interfaces for mass spectrometers are characteristically intolerant of salt content in the fluid stream. Development of a successful salt resistant or salt remediating interface would open downstream processing to the power of MS. This would appear to be the primary and only limiting factor in successful, robust application of MS in biotech process applications, notwithstanding the high acquisition and operating cost of a mass spectrometer. 

Liquid chromatography has been successfully employed in pharmaceutical PAT applications for decades. Of particular note are the pioneering examples of on-line high performance liquid chromatography (HPLC) applications by Eli Lilly in biotechnology applications.

  On-line downstream applications of HPLC for biotechnology are now widely accepted and deployed. Commercial systems for biotech process monitoring are available from Dionex and other suppliers. This technique is mature, well characterized, and well accepted in all fields of chemistry separation including biotechnology. The system also requires capture of a physical sample from the process stream while maintaining sterility of the stream. Recent advances in on-line sampling for biotechnology, including introduction of commercial products that maintain process sterility, will further increase utilization of the HPLC technique in downstream applications.

  The chief drawback of this technique for process characterization appears to be the complexity of fully automated HPLC systems and their attendant control processes. These include column switching, fast column equilibration, and high flow systems, all of which are strategies employed to reduce cycle times to remain consistent with downstream needs.   

Electrophoretic techniques, most notably slab gel electrophoresis for protein and genetics analysis, have been a mainstay of biotechnology development analytical laboratories since its commercial inception in the 1970s. Slab gel electrophoresis is simple in concept, inexpensive to acquire, does have high operating costs, and has cycle times inconsistent with the needs of downstream processing. Capillary electrophoresis (HPCE), was introduced in the late 1980s and automates the electrophoresis process in a format similar to HPLC. Many commercial instruments are available today. Data are quantitative in contrast to slab gel techniques. It is rapid. It has recently found widespread application for protein analysis in off line laboratories operated in support of downstream applications, specifically for protein product quality, characterization, and quantitation. The technique is inherently fast, versatile in the biotechnology environment, and amenable to the demands of protein assay in the downstream process environment. The technique is employed daily in this capacity, but still has failed to gain wider appreciation and acceptance for downstream processing assay requirements because commercial instruments have been designed exclusively for standard benchtop laboratory use. Manual sample acquisition, preparation, sample introduction, and data reduction are the rule as is the fact that most HPCE systems must be operated by HPCE specialists. These instruments have remained exclusively in the province of remote laboratories operating in support of downstream operations and these instruments do not promote the advantages of PAT technology in downstream operations. 

We are introducing an on-line and at-line process analysis technology specific for protein identification and quantitation in a downstream process application. This instrument

  fully automates the entire process and is purposely built for on-line assays for protein identity and titer as envisioned by the FDA PAT initiative. The instrument incorporates and can be used on-line with an automatic, sanitary sample port interface, or can be used at-line with samples introduced by the instrument's autosampler. In either case, the entire sample introduction, preparation, separation, detection, and data reduction are performed automatically by the instrument without operator input. Once the sample is introduced to the instrument it is transferred directly to the sample preparation module, which delivers an appropriately prepared sample to a HPCE module that automatically performs the separation. Detection is performed by an ultraviolet absorbance detector. Data reduction and reporting are performed by a data system based on standard HPLC data and user interfaces. The concentration dynamic range of the instrument is from ?g/mL to g/mL due to the internal automatic dilution-concentration system component of the sample preparation module. Molecular weight range for the instrument is 10 to 200 kD. Calibration of the instrument is through a timed, automatic introduction of calibration standards in a manner typical of calibration for similar HPLC systems. Cycle times are consistent with the needs for most downstream processing. 

Biotechnology manufacturing, particularly downstream processes, has many operations and opportunities suited to the premise of the PAT initiative—namely, improving production efficiency, yield, and product quality. Several technologies have been implemented in downstream processing for the purpose of on-line assays that have proven the concept. New, versatile instrumentation technologies for downstream process applications will continue to emerge as the PAT initiative becomes more fully implemented in biotechnology and biological manufacturing. 

 is president and CSO of Groton Biosystems, 85 Swanson Road, Boxborough, MA 01719, 987.266.9222, fax 978.266.9223		

 2. PAT definition per FDA CDER. A system for designing, analyzing, and controlling manufacturing through timely measurements (during processing) of critical quality and performance attributes of raw and in-process materials and processes with the goal of ensuring final product quality. It is important to note that the term analytical in PAT is viewed broadly to include chemical, physical, microbiological, mathematical, and risk analysis conducted in an integrated manner. 

 3. Sharaf MA, Illman DL, Kowalski B. Chemometrics. John Wiley and Sons; 1986. 

 4. Brereton RG. Chemometrics: Data Analysis for the Laboratory and Chemical Plant. John Wiley and Sons; 2003. 

 5. Brereton RG. Chemometrics and PAT. PAT Journal. 2005 2 (3) 8-11. 

 6. Willis RC. PAT Pending. Modern Drug Discovery. 2004 December. 

 7. Cooley RE. Utilizing PAT to Monitor and Control Bulk Technology Processes. Eli Lilly and Co. University of Michigan Pharmaceutical Engineering Seminars. 2003 March 4 . 

 8. On-line sampling systems for biotechnology applications are supplied by Groton Biosystems, YSI Instruments, and Nova Biosystems, among others. 

 9. Groton Biosystems GPA1000 Process Analyzer. 

The purpose of the PAT initiative is to move analytical laboratory functions close to the manufacturing process to improve manufacturing efficiency and product quality.

Downstream Process Optimization Opportunities Using On-Line and At-Line PAT Instrumentation

Biotech companies are running into production bottlenecks because standard purification and separation technologies lack the capability to remove the elevated levels of biomass from high titer solutions. Recent developments in filter technology offer the biotech industry a cost-effective solution to processing challenges by reducing bottlenecks, thereby accelerating the time-to-market of new drugs. 

This article focuses on new designs of sterilizing-grade filters that remove bacteria, toxins, and other contaminants from initial purification steps to final formulation and filling. We discuss these important parameters in filter selection: materials, compatibility, wettability, sterilization, adsorption, and membrane and filter construction. 			

Three hydrophilic sterilizing-grade membrane materials are readily available: Nylon 6,6, polyvinylidene fluoride (PVDF), and polyether sulfone (PES). Table 1 summarizes the general compatibilities for each membrane type. PES and PVDF are low binding, water wettable and can be gamma irradiated. The membrane with the broadest pH compatibility is PES. 

 Table 1. General compatibility of polymers for filtration			

The ability to wet a filter completely is important, because the objective is to use the full filtration area. If a significant portion of the membrane is not wetted properly, then the filtration flow can be impaired. In addition, it is necessary to have a fully wetted membrane during the performance of an integrity test. If the filter membrane is not completely wetted, then even a very small non-wetted area of the filter could cause a failure of the integrity test, causing process delays and re-testing.  

In terms of wettability of membrane material, Nylon 6,6 is the most easily wetted because it is naturally hydrophilic. PVDF is naturally hydrophobic, and for liquid applications will under go some sort of modification to make the membrane hydrophilic. This version is close to Nylon 6,6 in terms of wettability. Most common PES filters do not wet easily and usually require backpressure to wet completely, making integrity testing more difficult and time consuming. However, recent improvements in PES wetting chemistry have rendered its wettability comparable to PVDF membrane filters.  

Integrated single-use bioprocess systems incorporating filters, tubing, and bags can play a key role in helping companies to increase manufacturing capacity. A single-use capsule does not require cleaning or cleaning validation because it can be pre-sterilized via gamma irradiation by the manufacturer. This simplifies process operations because pre-sterilized filters do not require assembly or steam sterilization. The housing material used in capsule filters is also often polypropylene. For capsules that will be sterilized by gamma irradiation, the polypropylene must be gamma stable to ensure structural strength, compatibility, and low extractables. 

Figure 1. This diagram depicts an asymmetric membrane with a 3:1 gradient in average pore size			

The purpose of filtration is to remove a particular contaminant or groups of contaminants from the process feed. The filter system should not be adding anything significant (extractables) to the process fluid, nor should it remove a desired product component from the fluid that is being filtered.   

In some cases, where low concentrations of proteins or preservatives are in the fluid, there may be a loss in product yield if the filter membrane adsorbs significant amonts of product . At high component concentrations this is not a concern, but this is especially critical where a desired component is present in very low concentration. The effluent should be tested to ensure that the amount of loss is acceptable. 

Here is a test run example. IgG protein transmission was measured to demonstrate differences in protein adsorption among membrane types. Each disc was challenged with 60 mL protein solution at 100 μg/mL concentration. Filtrate was collected in 1 mL aliquots for the first 10 mL followed by 5 mL aliquots to a total throughput of 60 mL. Data in Table 2 indicate that protein binding occurs within the first 10 mL passed through each membrane disc. After the first 10 mL, the effluent contains protein at the same concentration as the influent solution. PES and PVDF are generally low adsorptive materials. However, nylon bound 20 times more protein than PES in the first 10 mL. Extrapolating to production scale; this may affect product yield. 

New filter designs have faster flow rates, smaller filter systems, improved microorganism and particle retention, and longer service life. These benefits translate into significant cost and time savings for drug manufacturers. Advances in the speed, size, and performance of new generation membrane filters are perhaps most critical because membrane technology is used in a broad range of applications. Lifetime (throughput) and flowrate at process scale are highly influenced by the structure of the membrane.  

When flowrate is the main driver, new high-flow sterilizing-grade filters feature a serial layer construction where the upstream layer is an asymmetric, high porosity PES membrane over the final sterilizing-grade layer. This PES filter offers exceptionally high flows at a processing rate of 17 L/min at 100-mbar differential pressure (916;P), and is compatible with fluids across a wide pH range, from buffers to numerous biological fluids. 

Figure 2. Cross section of high-area asymmetric filter cartridge showing retained contaminant			

When lifetime is the main driver, a high-capacity sterilizing-grade filter offers users an innovative hybrid technology by fitting a finer grade of asymmetric PES membrane over a PVDF membrane. This membrane combination enables high dirt-holding capacities and the flexibility for either wet or dry steaming. Because  autoclaving or steam-in-place procedures can take place in either wet or dry conditions, this flexibility accommodates different manufacturing specifications. The high-capacity hybrid filter offers competitive flow rates at 10 L/min at 100-mbar ΔP; in addition it offers the high throughput required in the filtration of high protein biotech solutions. 

Figure 3. Upstream side of prefilter final filter layers showing clean membrane due to effective prefiltration			

Asymmetric Prefilter Construction Improves Capacity

A technology that has contributed significantly to the performance of new generation membrane filters is an asymmetric  membrane structure that offers a 3 to 1 average pore size gradient through the depth of the membrane, resulting in higher porosity and dirt holding capacity. This pore size gradient (Figure 1) enables larger particles to be captured in the outer portion of the filter, while the sterilizing layer provides high efficiency removal of smaller microorganisms. 

Figure 4. A unique folding pattern in a new filter design increases filter area			

Whereas traditional membrane constructions have a homogeneous pore size distribution across the membrane, the asymmetric structure with its large void volume is able to capture solids across the entire depth and pore gradient of the membrane, improving total filter capacity. (Figures 2 and 3). 

Table 4. Cartridge construction affects flowrates			

Minimizing system size while maintaining process flow and volume has required unique approaches. One of these is modifying the traditional pleat pack design. In order to achieve greater filter surface area, while keeping the same cartridge  configuration the traditional fan pleat filter has been replaced with a deep pleat design (Figure 4). This deep pleating technology is used to form a very robust filter pack that allows up to 50% greater filtration area than traditional fan pleat and improved flow distribution in the membrane cartridge. Table 3 shows the benefits of deep pleat design in that there is more area in the PES and hybrid filters compared to traditionally pleated nylon and PVDF filters. 

Table 5. Test of filtering 100 L of vaccine			

Another factor employed to increase pleat length is a reduction in the size of the core-collecting chamber. This allows the flow path to extend into the space that would normally be reserved for the core, enabling much greater membrane flow and fluid throughput (Figure 5).  

The core chamber's materials of construction also influence the integrity and usability of the filter. The combination of a narrow polypropylene core and deep pleat design enhance mechanical stability and robustness during operation and steam sterilization. The strength of the design means that a higher differential pressure (1 bar vs. 300 mbar for most fan pleated filters) can be safely reached during steam in place operation without damaging the filter. This means that a 10-in. filter can be used to achieve the flow rates of a conventional 20-in. filter. For example, a sterilizing-grade deep-pleat filter with 1 m

  membrane can achieve flow rates of greater than 17 L/min. A traditional 10-in. filter with the same removal efficiency might only provide flow rates of 3 to 7 L/min. Figure 6 shows results of a test. By increasing flowrate capability of single filter elements, the deep pleat design also allows biopharmaceutical companies to reduce the number and size of filter assemblies. 

Filterability studies were performed using model buffer solutions to determine the optimum membrane for large-scale buffer filtration applications. For solutions with low solids levels such as clean buffer solutions, results are best described in terms of flowrate per 10-in. cartridge per hour, at a given pressure, rather than volume processed to point of plugging. Table 4 shows that the high-flow PES membrane demonstrated flow rates 3 to 5 times greater than the nylon membrane and 3 to 4 times greater than the PVDF membrane. Based on the results, PES membrane demonstrated superior flow characteristics during filtration. This is our membrane choice for buffer filtration applications. 

To filter a batch of 100 L of vaccine product a total of six traditional nylon filters would be used. The same batch can be filtered with one deep pleated PES filter (Table 5). The deep pleat design and asymmetric pore structure allows end users to reduce the number and size of filter assemblies. Smaller filter assemblies mean fewer components to sterilize, test, clean, and validate. 

Figure 6. Water flowrates per 10-inch cartridge			

To process a 550 L batch of monoclonal product, two high-flow PES filters are required. The same batch can be filtered with one high-capacity hybrid filter (Table 6). 

As these examples have shown, however a manufacturer chooses to realize the benefits of innovative filter technology, the end result is more efficient and faster processing of lifesaving drugs. 

is a marketing manager with Western Hemisphere responsibility for direct flow filtration with Pall Corporation, 516.801.9526,  fax 516.801.9548, 

is an R&D mechanical engineer at Pall Corporation Life Sciences Department, +44.23.9230.3300, Oliver G. 

Biotech companies are running into production bottlenecks because standard purification and separation technologies lack the capability to remove the elevated levels of biomass from high titer solutions. Recent developments in filter technology offer the biotech industry a cost-effective solution to processing challenges by reducing bottlenecks, thereby accelerating the time-to-market of new drugs.

Filtration Designs Remove Processing Bottlenecks for High-Yield Biotech Drugs

Developing biopharmaceutical manufacturing unit operations inevitably involves screening chromatographic adsorbents that offer high selectivity and yield highly purified materials. Simultaneously, materials must meet the diverse and rigorous requirements for use in an economically viable cGMP process.  

When considering the technical performance of chromatography media, the selection process might involve preparing numerous packed columns, followed by a target protein challenging various candidate adsorbents to allow scrutiny of protein recovery, yield, purity, and activity. Then the capacity, affinity, and selectivity of an adsorbent are considered in the context of process conditions such as binding and elution buffers, or response to repeated clean-in-place (CIP) procedures. 

Packed columns have been commonly used in such process development activities. Using packed columns, however,  limits the range of media that can be appraised because packing the media in small-scale columns is onerous and may be hindered by the limited availability of (sometimes costly) automated systems that will perform method development or scouting experiments. As a result, optimal adsorbents or even process conditions may not always be realized or implemented. 

However, using 96-well microplates containing chromatography media to facilitate high throughput experimentation may rapidly enhance adsorbent selection, thereby allowing a greater opportunity to identify an optimum adsorbent and set of operating conditions. An investigation can  facilitate the appraisal of a range of adsorbents in a single plate, or develop the process conditions for a single adsorbent by challenging it with various bind and elute conditions. 

 Table 1. Summary of dimensions and characteristics of 96-well microplates used for adsorbent selection and process development.			

In this article we discuss adapting the high throughput ex-perimental advantage of 96-well microplate technologies for process selection and development. We highlight various aspects that need to be considered in developing a robust and trustworthy system of adsorbent appraisal. 

Microtiter plates in a 96-well format have been used for many years in analytical research and clinical diagnostic laboratories because of their high throughput of samples. Examples of their application include the enzyme-linked immunosorbent assay (ELISA) and high-throughput screening of pharmaceutical candidates. The speed, accuracy, and efficiency of these activities is often further enhanced by using robotic systems which can dispense and aspirate relevant fluids into specified areas of the plate, and manage multiple plates simultaneously. 

Figure 1. Summary of anticipated and observed effects when ion exchangers are exposed to pH and salinity gradients.			

Our laboratories have been using robotics in combination with 96-well microplate formats for many years to screen libraries of ligand candidates, which, on identification of promising results, can be developed into adsorbents used in the full-scale manufacture of biopharmaceuticals. Using  96-well microplates significantly increases the number of ligand candidates that can be screened as potential adsorbents in a given time frame and correspondingly, the chance of project success and its quality may also be improved. In addition, the small quantity of media used in each well means that relatively small quantities of test material are required to screen a high number of adsorbents. These reduced quantity requirements are helpful because often, representative feed material is in short supply, particularly at the onset of biopharmaceutical development campaigns. 

Figure 2. E-PAGE analysis showing elution profiles from SP PuraBead HF ion exchange media. Proteins are labeled based on the profiles observed in the calibration part of the E-PAGE (lane 10). White circles represent elution conditions that were later verified using a conventional column.			

We made specially adapted micro-plates that have an outlet in the base beneath each well. These allow the passage of fluid through the well and the adsorbent, while a polypropylene frit in the base of the well retains the adsorbent within the well. The dimensions and characteristics of these adapted 96-well plates appear in Table 1.			

Figure 3. E-PAGE showing elution profiles from CM PuraBead HF ion exchange media.When these plates are used, known quantities of chromatography media are aliquoted into 64 of the wells, and the remaining 32 wells are reserved for calibrations, standards, and controls. In this way, flow-through, elution, and sanitization samples all can be generated for further analysis. 

We routinely apply various analytical techniques to qualify and quantify both target proteins and impurities that have been challenged by the multitude of candidate ligand libraries stored in the plates. Our most common combination is Bradford's Protein Assay and Invitrogen's E-PAGE 96 Protein Electrophoresis System. This way, binding proteins are identified and can be compared with non-binding proteins to pinpoint the purification capability of any adsorbent. A positive aspect about microplate format is that additional analytical procedures (such as ELISA)can be readily applied to obtain more information from any well. 

Using Plates for Media Selection and Process Development

We have been investigating the potential use of microplate formats in two key areas: media selection and process development. In order to develop this tool as a product rather than an in-house technique, we have made some minor modifications to the composite of the microplate, frits, and adsorbent of the plates that are used in-house. Specifically, adding an upper frit to the wells means that the chromatography media is not unduly disrupted during shipping. Controlling the extent to which this frit compresses the media enables control over the flowrate of fluids passing through the resultant mini-column. In effect, the amount of compression controls residence time, and we have developed protocols that allow both selection and good reproducibility of residence times in the region of 100 to 300 seconds. The upper range of these residence times reflects what might typically be used in a large process-scale column. 

Figure 4. E-PAGE showing elution profiles from Q PuraBead HF ion exchange media.			

In our procedure for process development, we filled 64 of the 96 wells with the same adsorbent and then explored optimum elution conditions for separating commonly encountered proteins, as detailed in Table 2. Four different plates were used to illustrate their capability in small-scale process development, containing ProMetic BioSciences Q PuraBead HF, DEAE PuraBead HF, SP PuraBead HF, and CM PuraBead HF ion exchange media, which we abbreviate as Q, DEAE, SP, CM respectively. Mock feedstocks were prepared in equilibration buffers as described in Table 3. These feedstocks are composed of two proteins that can be readily distinguished by E-PAGE, and challenged to plates under binding conditions. Figure 1 displays the trends in pH and salinity, moving down and across the microplate respectively, and also states the anticipated, differing responses from anion and cation exchange media. 

Figure 5. E-PAGE showing elution profiles from DEAE PuraBead HF ion exchange media.			

Total protein was measured using Bradford's Protein Assay at the following stages: flow-through, wash, elution, and sanitization. Significant quantities of protein were only observed in elution and sanitization fractions. After pH and salinity gradients were applied across the plate, elution profiles were also monitored using the E-PAGE 96 Protein Electrophoresis System as shown in summary in Figures 2 to 5. The white circles represent elution conditions that were later compared for verification using a conventional column format. These conditions are listed in Table 4. 

Table 2. Proteins used to develop elution strategies from ion-exchange media. Anion exchangers (Q, DEAE) were challenged with a mix of polyclonal IgG and BSA. Cation exchangers were challenged with a mix of lysozyme and polyclonal IgG.  			

Four 96-well plates were used to illustrate elution and separation of proteins. Each plate contained uniquely one ion-exchanger of these four: SP, CM, Q and DEAE, all on PuraBead HF 6% agarose. The results of eluting the bound protein from each well of each plate were illustrated by the E-PAGE system. The eluate from each well was transferred to the corresponding location in the gel, and scrutiny of these E-PAGE profiles readily identified the originating well. Therefore, conditions that were applied to facilitate separation of IgG and BSA (in the case of the anion exchangers) or IgG and lysozyme (in the case of the cation exchangers) could be determined by referring back to the pH and NaCl concentrations applied to that particular well. 

Figure 6. SDS PAGE of challenge and elution materials from ion exchangers. Lane 1 (SeeBlue standard); lanes 3â5 (SP Load, 1st elution, 2nd elution); lanes 7â9 (CM Load, 1st elution, 2nd elution); lanes 11â13 (Q Load, 1st elution, 2nd elution); and lanes 15â17 (DEAE Load, 1st elution, 2nd elution). Laboratory notebook reference BG/NB8/1.			

Microplate data do not necessarily represent a developed process; more assurance is required. We tested the optimum results from each plate in a traditional packed column. The elution conditions achieved to desorb a single protein, and then both proteins, as observed on the E-PAGE when relatively intense bands on the gel were seen, were applied to a traditional packed column, and run on an automated workstation. 

Table 3. Summary of buffers used to analyze protein binding and elution from microplates containing ion-exchange media. Sanitization was carried out using 0.5 M NaOH, and media were stored in 20% ethanol. The plate is depicted in Figure 1.			

The ion exchangers' parameters were scaled up to ensure good correlation of results between the 96-well chromatography microplate study and conventional column chromatography. Two elution conditions were chosen for each ion exchanger, based on the images from the E-PAGE experiment, which demonstrated the elution of a single protein and both proteins. The buffer conditions are summarized in Table 4. In all, eight experiments were performed (one for each elution condition) corresponding to the four 96-well microplate studies. The elution conditions applied for column chromatography are identified on the E-PAGE images by white circles. 

From observing SDS PAGE analysis for all four ion exchangers (Figure 6) it can be seen that the elution profiles from column chromatography indicate that only IgG is eluted from the column when presented with the first elution buffer (lanes 4, 8, 12, and 16), exactly as was observed for the 96-well chromatography microplate study. We see that a small quantity of BSA eluted from the Q-ion exchange media from the first elution buffer, and this might indicate a requirement to elute the IgG under slightly less stringent conditions. In lanes 5, 9, 13, and 17, both proteins are eluted. Again, this result correlates with the 96-well chromatography microplate study indicated by the white circles in Figures 2 to 5. This correlation indicates that the separation of proteins in column chromatography compares favorably with the separation of proteins developed under the same conditions in the 96-well chromatography microplate study. Thus, we have shown the validity of using a microplate-based approach for initial process development activities.  

Table 4. Optimum conditions for separating proteins as developed using microplates in conjunction with E-PAGE analysis.  White circles in Figures 2-5 reflect the chosen points, which were then applied to these conventional packed chromatography columns. 			

Figure 7.  Example composition and layout of 96-well microplate containing a range of adsorbents (Mimetic Ligand adsorbents, Prometic BioSciences) and two anion exchange media.The microplate format also can be used for media selection. Using the microplate format would offer re-searchers the ability to investigate the purification capabilities of numerous affinity adsorbents in short time frames with a minimum quantity of representative process material. In our laboratories, we have been able to acquire extensive and informative purification data from a single plate containing up to 64 adsorbents within two days.  

An example of a format (well layout) for this type of study is shown in Figure 7, which contains our complete range of Mimetic Adsorbents, and in this example, two anion exchange media. Depending on the nature of the target protein, plates with cation exchangers might be used. Each adsorbent is represented in four wells to provide greater confidence in any acquired data. Applying a representative feed stream across the plate followed by the analyses discussed earlier would readily point to any potential candidates that could be developed for use in a manufacturing situation. 			

Quick RecapUsing 96-well microplates for chromatography adsorbent selection and process development may expedite typical activities undertaken by process development groups. 

Using such a format requires smaller sample volumes and can rapidly generate data from a number of points than might otherwise be possible in the same time frame. As a result, this method is likely to be of benefit in this important area of biopharmaceutical process development. Furthermore, using these high throughput experimental techniques increases the number of adsorbents or process conditions that can be evaluated, potentially resulting in substantial time and cost savings in process development activities. 			

, 211 Cambridge Science Park, Cambridge, CB4 0ZA, UK, +44.1223.420300, fax +44.1223.433270, 

Using packed columns in process development activities limits the scope for appraising a large and diverse range of media.

Chromatography Process Development Using 96-Well Microplate Formats

Technology to manufacture high-dose therapeutic monoclonal antibodies (MAbs) at scale has evolved in several waves over the last two decades. Major cost reductions have been reported through the use of platform technologies upstream, downstream, and with the portfolio of analytical assays used.

  This review describes a reference downstream process developed at GE Healthcare

  and discusses technical and updated economical analysis of various process design options found in the literature and conference presentations. 

Antibody titres in mammalian cell culture have improved one hundred fold over the last 20 years, mainly through the ability to grow higher cell numbers (~10 fold) and through increasing productivity of those cells (~10 fold)

  (Figure 1). Recent reports from industry demonstrate that antibody titres higher than 1g/L are the rule, and 5 g/L readings are already on the horizon for processes currently under development. At the same time, manufacturers observe new challenges, resulting from the increased generation of antibody aggregates and increased presence of host-cell proteins from the high-producer cells used in such upgraded processes. 

Figure 1.Twenty years of development in mammalian cell culture have led to significant increases in product titres for monoclonal antibodies (MAbs), as a result of higher expression levels, faster process times, and higher cell densities. Factors contributing to these improvements have been seen in the media for growth and the production phase, feed composition and strategy, process control and design, and host-cell engineering.			

During the same period, purification by chromatography has improved its productivity by a factor of 10–20. This improvement was achieved mainly through the introduction of products allowing significantly shorter residence time at similar or higher binding capacity than previous generations of chromatography resins (Figure 2). Recent Protein A affinity resins and ion exchangers enable processing of 50 kg of antibody per batch per day using process schemes with a normal degree of optimization and without going to extremes in dimensioning the columns.

   In combination with up-to-date process engineering strategies that remove non-productive time from the critical path of manufacturing, these products are likely to resolve the near- to mid-term bottlenecking problem in the downstream process. 

 Figure 2. Twenty years of development in agarose resins have led to increased  stability, higher capacity, and better volumetric flow.			

It is important to realize, however, that significant improvement in process economy is still possible upstream, primarily by shortening the very slow cell culture process (2–3 weeks for harvesting)

  and by de-bottlenecking the link between cell culture and the relatively fast downstream process (1–5 day batch time, depending on degree of process intensification). Frequently, several (up to 5–6) large, staggered upstream lines are needed to feed one downstream production line. 

It is likely that the protein mass to be handled per batch will increase further with some "blockbuster" MAbs, but almost certainly not with all future MAb processes (i.e., for most promising medical indications). As a result, further improvements are desirable downstream of cell culture. End product purity and safety, as well as process robustness, are not debatable and cannot be compromised when considering alternate process design options or potential economic advantages. It is essential that alternatives under consideration be reviewed based on their practical value for today's process design work. 

It should also be noted that certain efficiency improvements in one stage of development may turn out to be costly or difficult to manage in later stages at full scale and with intensified processing. For example, using disposables is attractive for operating a multi-purpose plant and handling a large number of candidate MAbs in development, but is not necessarily more economical for regular, large-scale manufacturing. For certain situations, the savings created by using  disposables may be at risk due to process changes between clinical phases.  It is not easy to balance straightforward process economics against the advantages of an agile pilot plant.  

A number of generalizations are possible and a priority scheme for seeking economic improvements can be developed (Figure 3). Work on technical improvements should adhere to that order of priority to secure the greatest impact. 

Figure 3. Priority ranking of options to generate economic gains related to development and manufacturing			

The industry is communicating the success of using "platform technology" approaches to process development routines and to the ultimate process design.

   A platform consists of verified technology for cell biology, cell culture, downstream processing, and analytical work as used for a specific category of target active drug substance such as MAbs. Platforms are regularly updated with new, verified technology. It has been reported that using such platforms has supported time savings of several months on the critical path to clinical trials. Using platforms has allowed some companies to reduce the need for process alteration prior to full manufacturing scale-up. Gaining three months on the way into the clinic can potentially save tens of millions of US dollars (MUSD) in net present value (NPV) for companies looking at an average MAb (~500 MUSD peak annual sales).

  In other words, deviating from the platform concept or not developing a platform at all is one of the most costly strategy issues MAb- producing companies can face in relation to manufacturing process development. 

Many companies view the use of Protein A chromatography as a capture step essential to a successful downstream processing platform

 , and some results suggest that developing alternative capture steps, such as using small molecule ligands described as Protein A alternatives, may both take longer and result in reduced robustness.

   Just one failed batch as a result of a robustness issue would cost tens of millions of US dollars in lost sales value or a possible risk to the patient population. 

A key economic driver relates to the output from the production facility, i.e., the quantity of product produced in it per time. Facility output is primarily a function of its utilization (expressed as percent of total facility time used), the productivity of the cell culture (key parameters are cell culture time and product titres), productivity of the downstream process (main parameters are capacity to process product mass and volumes), and "smart" process engineering solutions (through the ability to remove unproductive time from the critical path delivering product to the filling plant). Depending on these factors, the cost per gram of antibody produced seems to vary between 100 and 1,000 USD/g (Figure 4). 

Figure 4. Shortening cell culture time and increasing facility use are two ways to reduce the cost of antibody produced. As seen in the table, reducing cell culture time from 14 to 10 days increases yearly production by 67 kg and reduces the cost per gram by 78 USD. The graph shows that increasing facility utilization from 20 to 100% can lead to a 5-fold reduction in the cost of goods.			

Downstream process design can increase facility output  through improved overall process yield or higher batch capacity in mass and volume. Higher yields enable the production of more drug product or a reduction in the number of batches required to satisfy the market need. Combining these changes could make room for production of additional products in the same facility. Use of recent technology as well as state-of-the-art process engineering will be necessary to achieve significant improvement in both of these categories. 

The single most effective improvement of drug manufacturing costs comes from resin and filter re-use optimization, in spite of cleaning and validation costs that need to be considered as a result. This view is particularly valid for the Protein A step, but also applies to ion exchangers in large-scale production with high batch frequency. A scenario comparing a  single-use membrane adsorber with re-use column chromatography was recently published.

  This paper discusses the cost comparison in great detail and includes all related costs in the evaluation, yet it uses an old ion exchange resin as the point of reference. Table 1 represents the main conclusion derived from this study as a cost comparison for 10 years of use together with a calculation based on a modern anion exchanger. Data were obtained following the same approach and using the raw data supplied in this comparison.

Table 1. Importance of using recent technology in cost of use estimates: The most recent technology offers the best long-term cost of use at slightly higher initial raw material investment combined with re-use and improved mass and volume capacity (from ref.9). 			

Other incremental improvements of a process step, either through optimization of process parameters or exchange of one resin for another or for a membrane, will only produce small cost reductions unless any of the following items can be achieved:  					

Overall process productivity improvements or time savings (satisfying higher product needs in an existing plant), 

Sizeable reallocation of facility capacity for other productive purposes, 

Reduced number of failed batches through improved robustness and lower production risk.

An example in line with the priority ranking shown in Figure 3 would be doubling cell culture titre and binding capacity for all steps in the purifications sequence, which could  cut in half the number of  batches required to meet market needs, thus releasing the remaining capacity for other manufacturing or alternatively to double the output of one product. Comparing capacities for first generation resins and recently introduced products makes it clear that such improvement is achievable when moving from old resins to new. A novel cation exchange resin has binding capacity of 80–120 g/L IgG at residence times of 2 to 6 min and typical pH and conductivity values for the feed stream. This represents twice the amount of useful capacity for SP Sepharose Fast Flow in a typical first generation process (Figures 2,5). 

Figure 5. Large-scale Chromaflow columns are typically used in the downstream processing of recombinant proteins. 			

The Protein A step (in good company with virus removal filtration) is often considered the most cost-intensive step in current platform downstream processes. Even this step, however, represents only about 3% of total manufacturing costs. Replacing Protein A  with a less costly resin will reduce the raw material cost, but will have little impact on the total product cost because of resulting productivity losses. There is a risk that such a small cost improvement will be accompanied by lower process performance of the new step; the benchmark would be tough: 95–97% yield and >99% purity. Similar constraints of improvement options would hold true for subsequent downstream steps. 

Thus, efforts to improve downstream technologies should focus more on the ability of the downstream process to handle an ever increasing mass of drug product rather than on the raw material or step costs. We believe that this interpretation will remain relevant until there is a paradigm shift to entirely different unit operations that offer similar overall purification performance, safety, and robustness without constraints in mass and volume handling and at a cost of ownership below current costs for membranes and resins. 

Figure 6. Schematic overview of consensus process design (left), resins for the generation 2 process (middle), and other options that have been tested by the industry (right).Initial data about the design of a reference process were recently published and are used for research purposes of GE Healthcare Life Sciences (Figure 6).

  This is a second generation process using similar purification principles and step sequences to the current, most widely used platform process used to manufacture today's MAbs as described in the literature and at recent scientific conferences. The current most popular process may use resin and membrane products from different suppliers, but almost all are from the same generation as Sepharose Fast Flow. Consequently, for the most part, the latest generation of downstream purification technology is not being used. 			

The purpose of the development project is to address challenges such as the increase in upstream productivity of monoclonal antibodies in an integrated fashion (not solely in selected step improvements), and to remove downstream process bottlenecks now and in the near future. Notwithstanding other development projects and their possible outcome, it is our goal that the resin technology selected for this process will remain relevant for most processes developed in the next 5–10 years and be able to remain in full scale production for the next two decades with no need to alter the fundamental design. This goal coincides with the need to avoid major process changes that delay development and increase regulatory costs in the industry. Improvements offered by the new reference process compared with current popular practices can be described by two categories: process robustness, and productivity and costs. 

In any downstream process sequence, the capture step dominates robustness in many ways. Capture transfers the drug product into a stable environment, concentrates the dilute feed ~10 fold, and ideally removes most impurities that could compromise the stability of the drug throughout the remainder of the process time. It also minimizes the level of impurities to a low level handled reproducibly by subsequent polishing steps, independent of feed stream quality variations. The robustness period can be extended to include robust performance between various development projects,  covering a wide range of monoclonal antibodies (i.e., a"platform"). 

Figure 7. The stability of a novel Protein A resin ("MAbSelect SuRe," GE Healthcare) following several hundred clean-in-place cycles with 0.1 M NaOH..			

A novel Protein A resin, MabSelect SuRe for second generation MAb capture, has been studied from various aspects directly related to robustness: stability of the matrix under cleaning in place (CIP) conditions (NaOH), stability of the ligand on exposure to feed, and interaction with various antibodies. The resin exhibits long-term stability to CIP regimes using sodium hydroxide (Figure 7) and the ligand offers protease stability significantly higher than the level of classical Protein A (Figure 8).

   While Protein A consists of five different subunits with different binding specificities, the novel ligand is comprised of four identical subunits. This aspect has been used to explain the observation of more homogeneous elution behavior for a wide selection of antibodies and Fc fusion proteins compared with a classic Protein A resin.

  The subsequent two chromatography steps have been successfully tested for their ability to remove remaining impurities to current acceptance levels.

Figure 8. The stability of the "SuRe" ligand (GE Healthcare, shown in red) and classical Protein A ligand (rPrA, shown in green) following incubation in CHO cell lysate for 18h at pH 5.0.			

Productivity and costs will vary between individual process designs developed by different groups testing the second generation process, depending on the specific antibody for which it is applied. Therefore, we have calculated a scenario with a 10,000-L fermentation volume containing 50 kg MAb (titre 5 g/L) using conservative assumptions for capacity and volumetric flow values comparing the first and second generation processes.

  The new reference process is designed to produce up to 50 kg of antibody per batch per day of processing time. The comparison indicates that it offers 45% lower operating costs calculated for downstream steps (harvesting to the final UF/DF step), and a reduction of time needed to process the MAb by 80% compared to the first generation process.

Process platforms described by different companies vary somewhat in the two post-Protein A steps. Compared with the design shown in Figure 7, some processes have a flow-through anion exchange step, usually with a Q ligand in position two. Others use hdrophobic interaction chromatography (HIC) or ceramic hydroxyapatite in one of the steps. In recent years, membrane adsorbers have been reported to be useful for the flow-through anion exchange step as an alternative to Q resin chromatography. Process step comparisons published for this step variant to date all use the first generation resins and are challenged by the second generation reference process described here (Table 1).

Modern downstream process technology allows processing of batch sizes equivalent to ton scale production per year. There is no reason to assume that future antibody drugs cannot be produced with technology that has been introduced in recent years. This finding is not a contradiction to the permanent need for improvements of process performance and economy. Solutions presented here may provide development teams with efficient options to manufacture MAbs and sufficient time to develop a third generation technology adaptable to manufacturing needs as they evolve in the future.  

Günter Jagschies, Anna Grönberg, Tomas Björkman, Karol Lacki, and Hans J. Johansson

  work at GE Healthcare Bio-Sciences AB, Uppsala Sweden, +46.18.6120880, fax +46.70.6121247, 

1.  Slaff G. Application of Technology Platforms to the Purification of Monoclonal Antibodies; presentation at BioProcess International European Conference, 2005 April 11–14, Berlin, Germany. 

2.  Grönberg A, Monié E, Murby M, Rodrigo G,Wallby E, Johansson HJ. A strategy for development of a mAb purification platform; poster presentation at BioProcess International European Conference, 2006 February 20–23, Prague, Czech Republic. 

Downstream process design can increase facility output through improved overall process yield or higher batch capacity in mass and volume.

Technical and Economical Evaluation of Downstream Processing Options for Monoclonal Antibody (Mab) Production

Severe Acute Respiratory Syndrome (SARS), a coronavirus (SARS CoV) is a respiratory disease, the main symptoms  of which include fever, cough, shortness of breath, and pneumonia. Because SARS has the potential to reappear as a new, naturally acquired outbreak or by accidental or intentional release, an effective vaccine is sought. One of the most promising candidates for a SARS vaccine is the S glycoprotein protein.  This glycoprotein forms large spikes in the viral envelope and mediates the binding of SARS CoV to the host cell through the host cell receptor, Angiotensin Converting Enzyme II (ACE2).

  A promising platform for the production of this SARS vaccine is insect cell cultures. Insect cells have the ability to produce proteins at high concentrations with translational modifications that permit proper folding and function. Additional advantages of insect cells over mammalian cells include the ease of culture, high molarity tolerance, and high expression levels. 

This increased demand for insect cell products leads, in turn, to greater need for effective harvesting methods for large-scale cultures. Techniques have been established for harvesting insect cells using batch centrifugation,

  and tangential flow filtration using either hollow fibers

  Batch centrifugation is the simplest method for small-scale cultures, but it is very difficult to scale up to larger sizes because each batch usually is limited to only a few liters. Continuous centrifugation permits larger scale operation, but requires considerable initial investment. Also, the concentrate contains many particles that are 1 Âµm or larger, which means an additional filtration step is necessary before further processing. Finally, the product that remains in the sediment-containing phase is not recovered.

An alternative to centrifugation is tangential flow filtration (TFF). TFF has the advantage of being linearly scalable from the bench top to large scale production.

  A major drawback to TFF is the tendency of microfiltration membranes to foul with particles in the media, which lowers membrane performance and makes cleaning difficult.

 Another complication of using TFF is the shear sensitivity of infected insect cells.

  If TFF is operated at too high a flow rate, the cells will break apart as a result of the shear. These cell parts further foul the membrane.  

This study tests the performance of a modified TFF system, "SmartFlow TFF" (SFTFF, NCSRT, Apex, NC), in obtaining a high yield of a target protein from insect cell culture. In SFTFF, ribs create uniform retentate channels with uniform flow patterns over the entire functional surface of the membrane module. These uniform flow patterns maximize membrane efficiency and minimize fouling of the membrane module. The uniform flow also makes it possible to scale the system linearly from the laboratory bench to commercial production. In installed systems for other applications, the system has been scaled up directly from 5-m

  laboratory studies to commercial operations that process process 1.4 million L/d.  

In this work, a single tank, single-module process was developed using SFTFF to harvest the SARS-CoV spike protein, which was expressed intracellularly from an insect cell culture. One microfiltration module was used for the following steps: (i) clarification; (ii) diafiltration to replace the medium with buffer; (iii) lysis of cells and release of the desired protein by the addition of extraction buffer; and (iv) passage of the released protein through the membrane. Then, an ultrafiltration membrane was used to concentrate the protein and perform a buffer exchange before loading the protein onto an HPLC column. No lysis was observed, which means that SFTFF did minimal harm to the cells during clarification. The one tank–one module method simplifies the harvest in comparison to multiple centrifugation steps. The implications of these results for both small-scale and large-scale harvesting of insect cells are discussed below.  

The gene encoding the S glycoprotein was cloned from a lysate of culture SARS-CoV 3200300841 (Passage #3) in Trizol LS Reagent (Sigma, St. Louis, MO), kindly provided by Dr. Dean Erdman (CDC, Atlanta, GA). The cDNA was cloned into a baculovirus transfer vector pSC12 (Protein Sciences Corporation, Meriden, CT) downstream from the baculovirus very late promoter of the polyhedron gene and in frame with the chitinase secretion signal sequence at the N-terminus and 6-his at the C terminus.  

To generate a recombinant baculovirus, linearized Autographa California Nuclear Polyhedrosis Virus (AcNPV) DNA, and the recombination plasmid DNA containing the S gene were mixed and co-precipitated with calcium phosphate, and Sf9 cells were transfected. Recombinant viruses were identified by their distinctive plaque morphology. A single recombinant virus plaque was isolated. The virus was further amplified using Sf9 cells for P1 and ExpresSF+ cells in the absence of serum for P2 and P3. The recombinant virus was further scaled up before added to 10 L of ExpresSF+ cell culture in a bioreactor at a multiplicity of infection of 1.0 plaque forming unit/cell. The infected culture was incubated at 27–308 C for 48 h. During this period, recombinant SARS-CoV FL–His-6 was not secreted and was associated intracellularly.  

A PUROSEP LT-2Q filtration skid equipped with an OPTISEP 3000 holder (NCSRT, Inc., Apex, NC) was used for all the TFF experiments. An OPTISEP 3000 module containing 0.186 m

  of a modified polysulphone membrane with a pore size of 0.8 Âµm and channel height of 0.5 mm was used for the clarification and protein passage. The protein concentration was performed using an OPTISEP 3000 module containing 0.093 m

The overall average flux rate for the concentration and diafiltration step was 41 L m–2 h–1.

BioPharm International-06-02-2006

The Renaissance of Protein Purification

Downstream Process Optimization Opportunities Using On-Line and At-Line PAT Instrumentation

Filtration Designs Remove Processing Bottlenecks for High-Yield Biotech Drugs

Chromatography Process Development Using 96-Well Microplate Formats

Technical and Economical Evaluation of Downstream Processing Options for Monoclonal Antibody (Mab) Production

Development of a Novel Platform TFF System for Insect Cell Culture Harvest