Increasing titers in mammalian cell culture has recently shifted the focus of process development efforts toward improving the economics of product recovery and purification processes. Some argue that a paradigm shift may be required to ensure timely and cost-effective delivery of future antibody candidates. This paper discusses the key process economic drivers and the impact of scale and titer on downstream processing (DSP) economic trade-offs.  

The success of therapeutic monoclonal antibodies (MAbs) in the treatment of indications such as cancer and autoimmune diseases has fuelled clinical trial activities; in recent years MAbs have become the fastest growing segment of the biopharmaceutical industry.

  Attractive returns coupled with the potential losses in revenue resulting from delays in product approval have made companies focus on speed-to-market rather than on improving process economics.

  However, MAbs are amongst the most expensive drugs—the annual cost per patient can reach $35,000 for antibodies that treat cancer conditions. With intensified competition on the way and increasing pressure from healthcare providers, the ultimate success of the next generation of MAb therapeutics will increasingly depend on economic factors.

  As a result, production costs and capacity use are becoming critical success factors for the industry.

The relative importance of key process parameters on the overall economic feasibility of a process varies with both the scale of operation and the titer assumed. As annual output and scale increase at a given titer or combined with titer increases, the relative importance of different cost categories are expected to change as follows (Figure 1):  

Overall cost of goods/gram (COG/g) decreases

 -Material costs rise considerably and dominate COG/g

 -Labor and capital-dependent costs (overheads) represent a less significant proportion of COG/g 

 -Downstream processing costs become a major component of COG/g.  

If the increase in annual output is also accompanied by titer increases, then the trends are expected to become even more pronounced. At small scales, fixed costs tend to dominate and any changes in raw materials will have minimal impact. However, as scale increases, the role of raw materials becomes more critical. It is therefore important to look at the distribution of the costs (upstream to downstream, and direct to indirect costs) and at the scale and titer of interest to prioritize optimization efforts. 

Efforts to lower COG/g must be targeted at decreasing the overall batch costs (e.g., reducing raw material costs) or increasing the overall productivity (e.g., increasing process yields). This section discusses specific downstream processing (DSP) drivers, which are of prime importance when handling multigram per liter titers.  

As titers increase to 10–15g/L, it is expected that this will have a profound effect on reducing COG/g, as long as the purification costs do not negate the cell culture gains. With increasing titers, the ratio of upstream to downstream costs shifts so that the downstream costs become more dominant. For example, Sommerfeld and Strube calculated that increasing the fermentation titer 10-fold from 0.1 to 1 g/L caused the ratio of upstream to downstream costs in their process to drop from 55:45 to 30:70. This shift reflects the fact that the upstream costs are inversely proportional to titer but the same is not true for the downstream processing costs.

  Increasing the titer to satisfy larger market demands increases the protein load on chromatography steps resulting in an increase in the number of cycles or additional investment in larger columns; this also produces larger volume loads on any subsequent filtration steps leading to longer filtration times or a need for larger areas.  

All these factors increase the downstream operating costs per batch. However, the overall COG/g can still fall if the increase in overall productivity outweighs the increase in downstream costs. Accordingly, as titers increase further, the downstream processing steps will become major contributors to the overall COG/g and offer greater potential for improvements and cost savings. Consequently, the downstream yield and material costs become significant cost drivers. DSP bottlenecks can lead to increased investment in larger equipment and longer batch durations. This results in increased running costs and decreased productivity and suboptimal COG/g values.  

The overall DSP yield is a function of the individual step yields and the number of downstream processing  steps, as has often been demonstrated using the plot in Figure 2. Improvements in step yields and the reductions in the number of steps have contributed to typical overall process yields increasing from 40 to 75% in recent years, with savings in cost of goods and investment, and allowing for higher facility throughputs.

The impact of increasing step yields has been illustrated by Sommerfeld and Strube where increasing the average step yield in a seven-step downstream process from 85 to 95%, which increases the overall yield from ~30 to 70%, results in a 40% reduction in the downstream COG/g.

  Increasing step yields actually increases the equipment size or number of cycles required and, hence, the cost of the DSP steps, because each step needs to handle a larger load (chromatography) or volume (filtration). However, because more product is produced per batch, the COG/g typically falls with increasing yields. 

To maximize productivity and minimize investment and running costs, it is advisable to keep the number of downstream processing steps to a minimum.

  For antibody processes, this has encouraged the elimination of buffer exchange steps (diafiltration) that add little purification value by designing each chromatography step so that it can take the material eluted from the previous step where possible.

  Some companies have recently adopted processes that use only two chromatography steps while maintaining the desired purity levels.

  This requires an anion-exchange step to have additional selectivity to replace the intermediate purification and polishing steps. In cases where the contaminant profile, pH, and conductivity provide the opportunity to adopt this simpler process, time and cost savings can be achieved.  

In downstream processing, the distribution of raw material costs is highly dependent on whether or not resins and filters are reused. When treating these material components as disposables, resins and filters tend to dominate the material costs. Similar patterns can be seen in clinical manufacturing because these materials remain product specific. The impact of this operating strategy on the overall COG/g can depend on both the scale and the phase of development. The use of downstream consumables such as resins and membranes in a disposable fashion for a 200-L antibody process supplying early-phase clinical trials can provide both financial and operational savings.

  It has been reported that downstream processing using disposables can become a major disadvantage at the 10,000-L scale.

  This can be attributed to the fact that economies of scale result in a disproportionate effect on raw materials.

  Consequently, raw materials savings become more important for any process as the scale increases. 

The reuse of resins and filters involves a trade-off between reduced material costs and increased cleaning validation costs to determine the number of reuses with consistent performance. The higher the component cost or number of process steps, or the lower the validation costs, the greater the incentives to adopt filter or resin reuse.

  Chromatography resins, in particular Protein A, are often quoted as dominating purification raw material costs, owing to the high cost of the resin which is higher than ion-exchange resins. Large bioreactor scales of 10,000 L operating with a titer of 1 g/L, can result in Protein A resin costs of $4–5 million.

  Consequently, Protein A resins tend to be used in smaller quantities with multiple cycles, despite the complications of reuse validation.

  The reuse of Protein A resins can dramatically reduce their relative contribution to costs, making the costs associated with filters much more prevalent. In particular, virus filtration can represent a large contributor to purification material costs because of the costly membranes that are often used in a disposable fashion.

As the reuse of filters and resins increases, the cost of made-up buffers [chemical reagents and water for injection (WFI)] can account for a surprisingly large proportion of the costs, which can, in some cases, be greater than the cost of resins and filters. For example, at a fermentation scale of 20,000 L, approximately 140,000 L of buffer is required.

  The difficulty in estimating buffer costs partly reflects the large differences in the estimated costs of WFI, with values of approximately $0.20/L suggested for in-house generation and $3/L for vendor or contract manufacturing organization charges.

  Buffer costs have been quoted as varying between $2/L and $12/L.

  Efforts to reduce the volume and number of buffers required can lead to savings, because this naturally occurs when moving from a three-step chromatography platform to a two-step one. 

As increasing titers demand the use of larger downstream equipment or additional cycles, the requirement for buffers and WFI will also increase. In existing facilities, this can present retrofit challenges if demands exceed the capacity of the buffer preparation suites and WFI storage tanks or the rate of WFI generation. The use of buffer concentrates and in-line buffer mixing can help to reduce the tank size and floor space required for buffers.

The greater the mass load during chromatography, the greater the likelihood that the practical capacity of chromatography columns will be exceeded where the current limit of column diameters is 2 m.

  Under these circumstances, multiple cycles may be required and there is a risk that the downstream processing time will exceed the bioreactor time. This will reduce the potential throughput of the facility and impact on the COG/g. These large columns can also pose installation challenges in existing facilities if there is insufficient floor space and if they cannot fit through doors.

Process Economic Trade-offs for DSP Bottlenecks

Current efforts to avoid downstream processing becoming a bottleneck when handling larger masses include intensifying existing processes by enhancing capacity and speed. Such approaches can mitigate the need for extra investment in equipment and improve the process economics. However, with each of these approaches there are trade-offs and uncertainties that need to be evaluated to assess the impact on overall process economics.  

Chromatography Resin Dynamic Binding Capacity

Increasing resin binding capacity reduces column size requirements with concomitant drops in resin volumes required and buffer consumption [for equilibration, washing, elution, regeneration, and clean-in-place (CIP)] per batch. Novel resins have capacities that are more than twice those of the first generation resins.

  This can lower consumables costs which are a major component of COG/g at higher titers and demands,

  although the impact on the consumable costs will depend on the price differential between first and second generation resins. Increasing the binding capacity of affinity resins can have a greater influence on COG/g because affinity resins are more expensive than ion-exchange resins. With the cheaper ion-exchange resins, Sommerfeld and Strube highlight that a trade-off exists between the less pronounced drop in consumables cost and the increase in labor costs, which becomes more important at higher binding capacities, because of the longer processing times. However, given that most of the novel resins also allow higher flow rates, this may not be an issue. 

The first generation of cell-culture–based MAb processes used compressible chromatography resins. However, these impose severe limits on usable bed heights and flow rates when considering the expected increases in titer and scale.

  A move away from compressible resins and towards rigid resins that can handle higher flow rates reduces process cycle times and turnaround time and increases productivity.

  Flow rates with rigid resins can be three to five times faster than conventional compressible agarose resins.

  However, their use can also lead to increased buffer demands, higher pressure drops, packing complications, and shear stresses.

  New ion-exchange resins have recently been developed that can handle high flow rates (700 cm/h) and low back pressures (<3 bar). If the productivity increases outweigh the increased buffer demands, this will positively influence the COG/g.  

As mentioned earlier, resin and filter re-use can have a significant impact on the process economics at higher scales if the materials are expensive, despite the CIP and cleaning validation costs. Re-use also reduces the frequency of column packing which is time-consuming and costly when carried out on a large-scale.

  New resins are becoming available with increased stability when exposed to the harsh chemicals used for CIP; hence, they have longer cycle limits and can contribute to reducing the raw material costs.  

Platform processes provide a generic approach to antibody production that greatly reduces the development time while streamlining the regulatory aspects of processing. They represent a useful starting point for customization depending on the antibody being manufactured. Advances in resin properties have also allowed platform processes to emerge with two rather than three chromatography steps.

  This can help to alleviate DSP bottlenecks in existing facilities because a two-chromatography–step process occupies less floor space and consumes less buffer. Through such process intensification methods, Kelley predicted that a platform consisting of two chromatography steps with high capacity resins would be able to handle an annual output of 10 tons.

  Newer resins with the combined attributes of longer lifetimes, higher flow rates, and improved dynamic binding capacities will lead to improved platform processes for antibodies and contribute to significant reductions in downstream costs.

Research into alternatives for column chromatography focuses on methods that have the potential to effectively handle increased amounts of both the product and impurities (e.g., host cell proteins and antibody aggregates or isomers). Ideally, these alternatives should achieve a separation power equal to that of column chromatography while reducing the COG/g.

  When assessing the cost-effectiveness of these alternatives, it is important to consider not only the equipment sizes and resource consumption, but also the development and validation costs required.  

Membrane chromatography operating in flow-through mode is emerging as a popular alternative to anion-exchange chromatography steps in MAb purification,because of its rapid operation, ease of scale-up, and cost savings (Table 1).

  The dominant component in the distribution of raw material costs shifts from buffer costs in packed-bed chromatography to membrane costs; a membrane suitable for processing a batch of several thousand liters can cost several thousands of dollars and is disposable and not reusable. The key process economic trade-offs for anion-exchange applications therefore depends on whether the savings in buffer, labor, and overheads outweigh the high cost of the membranes. Critical variables that will affect the outcome of this cost comparison are the relative differences in the handling capacities assumed between anion-exchange membranes and resins, which dictate the sizes required, and the assumed WFI and buffer costs; higher values of these variables increase the economic attractiveness of membrane chromatography.

  The pace at which resin and membrane capacities improve will contribute to which operation secures its place in future platform processes. In cases where packed-bed and membrane chromatography offer similar COG/g, the real cost advantages may be in the development and validation costs that are significantly reduced with membrane chromatography because there is no column packing or cleaning validation.

Table 1.  Example of downstream process economic trade-offs 

As demand and titers continue to increase for MAbs, the DSP costs will become an increasingly dominant proportion of the COG/g with the DSP handling capacity representing a potential bottleneck that could reduce productivity. These factors have encouraged a shift in development efforts toward new DSP solutions that improve the process economics and alleviate bottlenecks. Consequently, the industry is taking advantage of improvements that affect the critical process economic drivers by looking to: improve the overall DSP yield and reduce the batch duration using platform processes based on two chromatography operations without intermediate buffer exchange steps; increase DSP capacity by taking advantage of improvements in chromatography resins that allow increased throughput over shorter times; and lower buffer demands and validation costs using new technologies such as membrane chromatography. 

Articles

The future of therapeutic MAbs lies in the development of economically feasible downstream processes.

Economic Drivers and Trade-Offs in Antibody Purification Processes

Monoclonal antibody purification processes have been challenged recently to recover 5 g/L cell culture titers in existing manufacturing facilities. This requires the modification of a platform purification process comprising Protein A chromatography as the robust capture step followed by two ion exchange chromatography steps. Because buffer and pool tank volumes are the primary facility bottlenecks, process development efforts have focused primarily on the order of unit operations, the dynamic binding capacity of current and new resins, reducing pool manipulations and buffer consumption, and elution buffer selection. This article presents examples in which new operating conditions or purification technologies can help to accommodate increases in titer without extensive changes to the manufacturing facility or equipment requirements.

Over the past five years, there has been a dramatic increase in the titers of recombinant proteins, particularly in the production of IgG monoclonal antibodies (MAbs) using Chinese hamster ovary (CHO) cells. Improvements in cell lines, media composition, and cell culture operating conditions have all contributed to higher expression levels. In most cases, these changes have had little or no impact on manufacturing equipment and facilities. However, as titers have increased by an order of magnitude or more using existing cell culture bioreactors, there has been a concomitant increase in the starting mass of protein entering downstream purification. This development has resulted in a shift in emphasis from increasing volumetric productivity for low-titer processes to handling significant increases in the amount of protein during downstream processing, particularly in existing facilities. Although incremental increases in titers can be handled by increasing the scale of purification unit operations (e.g., using larger chromatography columns and filters), at some point linear scaling exceeds the physical limits of existing facilities.  

To address the purification challenges of high antibody titers, it is imperative to determine the specific limitations of a company's manufacturing facility or facilities. This can best be accomplished by using a model that performs a facility fit using an existing process. Such a model can identify the bottlenecks for each unit operation at a specific manufacturing facility as a function of antibody titer. In some cases, it may be necessary to use multiple model iterations to find a bottleneck, model a change to the process or facility, and then determine where the next bottleneck may be. For the purposes of this article, we assume that one cell culture batch is purified to produce one bulk. Alternative process options such as split batch processes may be considered, and these could potentially allow higher recoverable titers from one cell culture batch. 

Almost all current antibody purification platform processes use Protein A affinity chromatography as the capture step because it is robust, widely applicable, and  facilitates the implementation of a successful standard purification process.

  Figure 1 shows two platform purification processes that we have implemented. After cell and product separation during harvest operations, Protein A affinity chromatography is used as the initial capture step with low-pH viral inactivation in the Protein A pool. Depending on the platform, this is followed by either the cation exchange chromatography step in bind-and-elute mode

  or the anion exchange chromatography step in flow-through mode. The virus filtration operation is placed between the two ion exchange chromatography steps. After the third chromatography step, ultrafiltration–diafiltration is performed using tangential flow filtration (TFF) for final product formulation.  

Potential volume bottlenecks caused by increased titers are included for each unit operation of each process shown in Figure 1. Underneath each potential bottleneck are process options that should be considered for optimization to avoid facility bottlenecks, and those having the greatest impact are shown in red. We have found that the primary bottlenecks in existing facilities are buffer and pool tank volumes, whereas secondary bottlenecks include pump capacities or pipe diameters that limit flow rates. For these reasons, the use of larger chromatography columns or filter surface areas is not sufficient to handle higher titers. 

One approach that allows greater amounts of antibody to be purified without increasing column volume is to increase the binding capacity of the chromatography step. Increasing the load capacity means the overall amount of processed antibody can be increased without increasing column size or volume. Since buffer volumes are usually based on column volume, increasing the load mass without increasing the column volume requires no increase in buffer volumes. The same applies for elution conditions where the pool volume will only be affected slightly by overall protein mass when using a final optical density (OD) cut off, or not affected at all when using constant volume pooling criteria. 

For Protein A chromatography, Table 1 shows the increase in titers that can be tolerated in an existing facility without increasing the size of the chromatography column. At a dynamic binding capacity (DBC) of 15 g/L, a maximum titer of 1 g/L can be accommodated in the facility. If the DBC can be doubled to 30 g/L, then the maximum recoverable titer increases to 3 g/L and a DBC of 45 g/L accommodates titers of up to 5 g/L. An increase in DBC can be achieved in several ways: by lowering the flow rate at constant bed height, increasing the bed height at constant flow rate to increase residence time, and/or by changing to a different Protein A resin with higher capacity.

  In addition, increasing the maximum manufacturing load density from low breakthrough levels (0.5–2% DBC) to slightly higher levels (3–5% DBC) also enables the purification of >5 g/L titers and most likely has an insignificant impact on step yield. Differences in DBC resulting from packing or resin lot-to-lot variability are considered negligible.  

Table 1.  Maximum expression level of a MAb that can be purified as a function of Protein A dynamic binding capacity. This assumes no other modifications were made to the facility, equipment, or operating conditions. 

Protein A resins all demonstrate higher DBCs with decreasing flow rate, although the slope of the curve may vary from resin to resin. As shown with the DBC curve for resin A in Figure 2, the DBC can be increased from 17 g/L at 40 column volumes (CV)/h to 38 g/L by decreasing the flow rate to 10 CV/h, therefore doubling the amount of antibody recovered with no other process changes. Processing time and facility throughput need to be assessed to determine whether reducing the flow rate also reduces overall capacity.

  This may be addressed in part by reducing the flow rate only for the loading phase of the Protein A step and then increasing the flow rate for the other phases such as equilibration, wash, elution, and regeneration. Unpublished data from our laboratories indicates that step yield is not affected by operating these other phases at higher flow rates. Another potential strategy is to conduct the load phase at two flow rates, and decrease the flow rate during the final stages of the load phase.

Table 2 shows how increasing the DBC affects a cation exchange chromatography step for a particular antibody and highlights the bottlenecks in an existing facility. In this example, a DBC of 50 g/L can accommodate a 1 g/L titer without constraints. A titer of 2 g/L is constrained by the buffer tank volume (the equilibration buffer in this case), and if this limitation is removed (perhaps by in-line dilution), the constraint becomes the pool tank volume at a titer of 3 g/L. Increasing the binding capacity of the cation exchange resin to 75 g/L or 100 g/L allows titers of 3 g/L and 4 g/L to be accommodated, respectively. In both cases, the buffer tank limitation occurs at lower titers than the pool tank volume limitation. This assumes that all other process parameters remain unchanged when the capacity increases. However, one key consideration during process development is the separation of the antibody and impurities at these higher load densities or with different (higher capacity) resins that may have different selectivities.  

Table 2.  Maximum expression level of a MAb that can be purified as a function of the dynamic binding capacity of cation exchange chromatography. This assumes no other modifications were made to the facility, equipment, or operating conditions. 

Decreasing Buffer Consumption and Using In-Line Dilution

As discussed above, it is important to take a holistic view of the process and carry out multiple model iterations that allow bottlenecks to be identified and removed, and subsequent bottlenecks to be identified. In examining the entire purification process, Table 3 shows the maximum titer that can be purified downstream by each unit operation. The baseline Protein A step can accommodate a titer of 1.3 g/L, but modification of the equilibration and various wash phases to minimize the Protein A buffer volumes increases this to 1.6 g/L. Table 4 shows an example of one equilibration phase and three separate wash phases in each Protein A chromatography cycle, where wash 1 and wash 3 also consist of equilibration buffer. In the baseline process, five column volumes are used for equilibration, three for wash 1, and three for wash 3. Studies have shown that this can be reduced to three column volumes for equilibration, two for wash 1, and two for wash 3 without affecting yield, purity, or column re-use, resulting in an overall reduction of four column volumes of equilibration buffer per cycle. For example, for a 140-L column operating eight cycles to process each cell culture batch, the revised process would reduce the consumption of equilibration buffer by 4,480 L per batch.  

Table 3.  Maximum expression level of a MAb that can be purified based on different process or facility modifications 

Since buffer tank volumes are a significant constraint throughout the purification process as antibody mass increases, one solution is to use concentrated buffers in volume-constrained tanks and then dilute each buffer to the desired concentration in-line to the chromatography column. This method uses various types of equipment (pumps, metering devices, and pH and conductivity sensors) to blend a concentrated buffer with water to achieve the desired concentration, which has been shown to attain the target concentration ±2%.

  The primary limitations of in-line buffer dilution are the ability to concentrate buffers sufficiently to fit into available tank volumes and the installation and validation of the in-line dilution equipment. As shown in Table 3, the maximum titer accommodated in the Protein A step increases from 1.3 g/L in the baseline case to 1.9 g/L using optimized phase durations and a 1.25x in-line buffer concentration factor. Using 2x concentrated buffers for the Protein A step would increase the capacity still further, accommodating titers of up to 2.9 g/L. The cation and anion exchange chromatography steps are also able to process more antibody through the use of in-line dilution and large column volumes. Although this approach requires capital expenditure and plant down time for installation, in-line dilution enhances the capacity and flexibility of a manufacturing facility. One additional benefit may be the option to move away from using fixed, stainless steel tanks for buffer storage and move to concentrated buffers in disposable bag systems.  

Table 4.  Operational sequence of a Protein A chromatography step and example phase durations in units of column volume (CV). 

Once buffer tank volume constraints have been addressed, pool tank volumes can become the next significant bottleneck. Pool tank constraints can arise during a given step through two mechanisms: a) the volume of the elution pool exceeds the tank capacity, or b) the pH and/or conductivity adjustments to condition the pool in preparation for the next chromatography step exceed the tank capacity. Increasing the resin DBC can address the first limitation, but the second limitation requires downstream unit operations to be investigated. Depending on the elution conditions and the target conditions for the subsequent step, a large volume of water may be necessary to lower the conductivity to the desired level for optimal antibody binding or impurity removal.  

Cation Exchange Process Optimization for Facility Fit (Process A)

When the cation exchange chromatography step is placed after the Protein A step, specific development considerations are needed to ensure both acceptable process performance and pool tank fit at different facilities. In some of our facilities, the bottleneck does not lie in the cation exchange pool tank itself, but rather in the pool tanks of the subsequent viral filtration and anion exchange steps. The cation exchange elution phase must be optimized to remove host cell protein (HCP), leached Protein A, and aggregates, while also minimizing pool volume and downstream pool conductivity. The anion exchange step functions to clear HCP and often requires a low load conductivity for acceptable process performance. The available operating space is often defined by a combination of factors affecting purity, product quality, product stability, yield, and manufacturing facility fit, as shown in Figure 3 for a cation exchange process. In this example, facility fit sets the upper limit for salt concentration, while both yield and purity are constrained by the interactive combination of elution buffer salt concentration and pH. One approach for debottlenecking the plant fit is to decrease downstream pool conductivities by eluting the cation exchange step at higher pH, thus lowering the salt concentration, assuming no negative impact on product quality at higher pH conditions.  

Newer anion exchange resins on the market have been designed for improved process performance at higher load conductivities. Table 5 compares two anion exchange resins, Q Sepharose Fast Flow (QSFF, GE Healthcare, Uppsala, Sweden), which has been a standard in antibody purification for some time, and Capto adhere (GE Healthcare), a new multimodal anion exchange resin introduced in 2007. In the flow-through process modeled in the platform process A example, QSFF requires a conductivity of 11 mS/cm or less to achieve optimal yield and requires 1,700 mL of water for each liter of the input pool to achieve this load conductivity. In contrast, Capto adhere was designed to have an optimal operation window at higher conductivities than previous anion exchange resins. At a load conductivity equal to that of QSFF (11 mS/cm), Capto adhere has a significantly poorer yield, but as the conductivity increases, the yield improves and is equal to that of QSFF at a conductivity of 22 mS/cm. The benefit to plant fit is the reduced volume of water (from 1,700 mL/L to 0 mL/L) needed to condition the load conductivity, removing the bottleneck caused by the pool tank volume limitation. As an alternative, TFF can be used to diafilter the pool, achieving the desired pH and lower conductivity, but this involves adding a new unit operation to the process. In addition, this increases the capital costs of installation, plant down time, and running costs. There must also be space available to install a TFF system in the existing plant.  

Table 5.  Impact of water required to adjust the conductivity for loading on an anion exchange chromatography resin used in platform process A 

Protein A Process Optimization for Facility Fit (Process B)

The Protein A process plays an important role in purification when the anion exchange chromatography step is next, as in platform process B. Protein A process parameters such as load density, elution buffer, and pooling criteria can have a significant effect on downstream pool volumes. The pH of the Protein A pool must meet low pH criteria for viral inactivation (pH ≤3.6),

  ideally without requiring additional acid titration. Process optimization must focus on minimizing the number of cycles, the pool volume per cycle, the volume of titrant required for pool pH adjustment, and the adjusted pool conductivity. Although maximizing the Protein A load density reduces the number of cycles, modifying the elution buffer components to minimize the pH and conductivity adjustment before the next chromatography step presents another alternative to address pool tank limitations. Figure 4 shows the impact of four different Protein A elution buffers on volume of titrant (Tris base) required to titrate the Protein A pool to pH 7.5 for loading onto the anion exchange step, and the resulting pool conductivity. There is a considerable difference in the amount of Tris base needed to adjust the pool pH to 7.5, ranging from 7 to 54 mL of Tris base per L of pool, depending on the Protein A elution buffer. Even more significantly, the acetic acid elution buffer resulted in the highest pool conductivity (~7 mS/cm), whereas the two glycine-based elution buffers resulted in pool conductivities close to 3 mS/cm. The lower conductivity load for the anion exchange step resulted in a reduction of HCP from 800 ng/mg to <5 ng/mg in the anion exchange pool for the two glycine buffers but only to 40 and 620 ng/mg, respectively, for the acetic acid and citrate elution buffers. Therefore, the glycine-based buffers offer the potential for a two-chromatography step process while the citrate buffer was eliminated because of its interference with the anion exchange process.

The need to model the entire downstream process to understand the effects on the performance of each unit operation is illustrated in Table 6. These results show that there is a slight impact on the overall pool volume for the final adjusted Protein A pool (~3,300 L compared to ~3,000 L) and that this increase is not constrained by available tank volumes. Because the anion exchange step is operated in the flow-through mode, there is an overall increase in volume in the anion exchange pool tank with all conditions resulting in ~4,000 L, which again is not constrained by tank capacity. However, due to the higher ionic components present in the acetic acid elution pool, a significantly large volume of water is needed to adjust this pool (450 mL/L) for loading onto the subsequent cation exchange step compared to either of the glycine-based elution buffer pools where no conductivity adjustment is needed. This may represent a bottleneck for a manufacturing facility because the adjusted anion exchange pool requires a tank with twice the capacity. Therefore, the glycine-based buffers deliver a robust process with lower impurity levels while also removing downstream pool tank bottlenecks by reducing pH titrant volumes and pool conductivities.  

Table 6.  Impact of Protein A elution buffer in platform process B on conditioning operations required for downstream chromatography operations.  Facility bottleneck is the load pool to the cation exchange step (marked in red). 

The optimization of operating conditions in platform purification processes allows titers of up to 5 g/L to be accommodated in existing manufacturing facilities. We and others have shown that downstream bottlenecks can be eliminated through the use of conventional purification technologies and the optimization of minor facility equipment.

  These platform processes use conventional technologies, including affinity and ion exchange chromatography, without the need for novel unit operations as previously discussed.

  Chromatography and other standard unit operations offer the benefit of consistent process robustness, high productivity, facilitation of platform processes, low raw material costs compared to the overall cost of manufacturing, and regulatory convenience. Since each antibody may bring its own unique challenges to a platform process, such processes should be flexible where needed while still permitting the use of standard conditions wherever possible. 

The authors would like to acknowledge Chris Dowd and Nuno Fontes for developing the facility fit model used in sections of this article and Jean Bender for discussions on process fit at multiple manufacturing facilities. 

is a senior research associate, all in late-stage purification; 

 is a scientist, early-stage purification, BRIAN KELLEY, PhD, is vice president, bioprocess development, and 

 is the director of late-stage purification, all at Genentech, Inc., South San Francisco, CA, 650.225.1956, 

 1. Jagschies G, Grönberg A, Björkman T, Lacki K, Johansson HJ. Technical and economical evaluation of downstream processing options for monoclonal antibody (Mab) production. BioPharm Int. 2006 June (supplement); 10–19. 

 2. Shukla A, Hubbard B, Tressel T, Guhan S, Low D. Downstream processing of monoclonal antibodies—Application of platform approaches. J Chromatogr B. 2007;848:28–39. 

 3. Fahrner RL, Knudsen HL, Basey CD, Galan W, Feuerhelm D, Vanderlaan M, Blank GS. Industrial purification of pharmaceutical antibodies: Development, operation, and validation of chromatography processes. Biotechnol Genet Eng Rev. 2001;18:301–27. 

 4. Swinnen K, Krul A, Van Goidsenhoven I, Van Tichelt N, Roosen A, Van Houdt K. Performance comparison of protein A affinity resins for the purification of monoclonal antibodies. J Chromatogr B. 2007;848:97–107. 

Altering the order of operations, using new resins, and increasing dynamic binding capacity can obviate the need for major facilty changes.

Purification Strategies to Process 5 g/L Titers of Monoclonal Antibodies

The following article describes the development of a purification platform for monoclonal antibodies (MAbs) at Pfizer's Global Biologics division, including how the individual unit operations have been affected by increasing cell culture titers and subsequent changes in the impurity profile, as well as viral clearance issues. The platform was developed to maximize the efficiency of the purification process and product purity while minimizing the development time for early-phase therapeutic antibodies. The success of this platform in its various stages has been demonstrated for several MAbs representing different IgG classes produced in different cell lines.

Recent advances in mammalian cell culture technology have resulted in significant increases in upstream productivity, with antibody titers of >5 g/L becoming increasingly common. Such increases could change the characteristics of the cell culture broth, leading to higher levels of product-related heterogeneous proteins (aggregates, degradation products, processing variants) and cell-derived impurities such as host cell protein (HCP) and cellular DNA. In response, platform purification processes for MAbs, i.e., those designed to move a product through Phase 1 clinical trials with minimum development, have become more streamlined in their design and execution. Such platforms must accommodate the majority of antibodies and yet be flexible enough to cover most contingencies and cell culture or antibody-related processing issues. They must also save time and thus accelerate the progress of antibodies toward the clinic.

The MAb purification platform used by Pfizer for early-phase clinical supplies has evolved over the last five years to accommodate increases in titer and associated issues, as well as unique challenges directly related to our own cell lines and upstream processes. This purification platform has been developed to accommodate antibodies derived from both NS0 and Chinese hamster ovary (CHO) cell lines. The original process, shown in Figure 1, followed a traditional antibody purification scheme.

  It consisted of three major chromatography steps with two additional unit operations designed to remove or inactivate viruses. After harvest, the cells were removed from the cell culture broth by depth filtration. In some cases, the broth was concentrated before the first chromatography step because of facility fit issues. Protein A chromatography was used in the initial capture step, where the antibody in cell culture broth was loaded at neutral pH, washed with various solutions, and eluted at a lower pH value. The eluate was then held at this low pH to inactivate viruses, and then adjusted to increase the pH before the cation exchange chromatography step.

After cation exchange, diafiltration was performed to prepare the product for anion exchange chromatography in flow through mode, in which impurities such as DNA and some HCPs were captured on the resin (and later removed by regeneration and cleaning steps). After anion exchange, the flow-through fraction containing the antibody was passed through a nanofiltration unit to remove potential viruses and then formulated in buffer in a final concentration and diafiltration step.

The relative purification development timeline for this original process is shown in Figure 2. Initially, several range-finding experiments were carried out to determine the optimal parameters for each unit operation and to maximize the yield and purity for that particular antibody. We used these results to determine the purification scheme, which was performed in its entirety at the bench scale before a larger demonstration run. After the demonstration run, this process was transferred to the pilot plant for manufacturing clinical supplies.

The original process was developed at Pfizer for a 500-L scale bioreactor. To accommodate an expanding pipeline, the scale of clinical manufacture was increased to 1,200 L, enabling the production of sufficient supplies in a shorter campaign. This increase in scale made it necessary to introduce changes into the platform process. One change reflected the efficiency of the initial depth filtration operation to remove cells and debris. During scale-up, we noted that effective filtration required a much larger membrane area than was feasible, so we introduced a continuous flow centrifugation process, followed by depth filtration to minimize membrane area, reduce costs, and increase the efficiency of cell removal.

To increase process efficiency, we also changed the order of the polishing chromatography steps, allowing the intermediate diafiltration step to be removed and thus reducing the total number of unit operations. Using this process, the pH of the Protein A affinity chromatography eluate was adjusted upward after pH inactivation and loaded directly onto the anion exchange resin. The anion exchange flow-through pool was then adjusted down to an optimal pH for the subsequent cation exchange step. All other unit operations remained the same. The resulting process is shown in Figure 1 as process 2.

In the next iteration of the process, further modifications were made to process 2 to reduce the clinical manufacturing time. The cation exchange polishing step was removed and the anion exchange chromatography step was changed from a resin-based operation to one involving membrane adsorbers. This increased the loading capacity of this step by a factor of up to 100.  All other unit operations remained the same. The resulting process is shown in Figure 1 as process 3. 

Then, the timeline for processes 2 and 3 was reduced by 30% (Figure 2). This was accomplished by performing previously defined experiments for each unit operation with specified ranges for buffer systems, load capacities, and pH/conductivity values. These experiments allowed ~80% of the process to be determined, with the remainder defined through additional bench-scale experiments specific to the antibody product. Subsequently, the experimental conditions promoting maximum yield and product purity were used to establish the parameters for each unit operation in the platform process. After the purification scheme was established, a larger demonstration run was carried out to evaluate scale-up effects before transferring the process to the pilot plant for manufacturing clinical supplies. 

Although process 3 worked well for many Pfizer antibodies, the performance was not satisfactory for all antibodies, particularly those produced at high titers with correspondingly high levels of impurities. In these cases, there were occasional issues surrounding virus clearance, raising safety concerns especially for antibodies required in large doses. To address these issues and increase the robustness of the platform, a cation exchange chromatography step was added to process 3 to define the current process (Figure 1). Based on knowledge gained from the implementation of the original process, and modified processes 2 and 3, the load capacities, buffer solutions, and other operational parameters were standardized to maximize purification efficiency for the majority of the antibodies in Pfizer's pipeline.

These changes reduced the timelines for purification development by 50% compared to the original process. The defined, optimized  platform process has been tested at the bench scale to verify that it is suitable for the production of clinical material. At this point, it must meet yield and purity specifications before it is scaled up for the demonstration run, or transferred to the pilot plant. This process is designed to meet the purification needs for 80% of the antibody products in our pipeline, which means the majority of the purification processes can be verified without further development. The remaining antibody projects may need slight modifications to meet individual specifications. These modifications will be evaluated on a case-by-case basis. 

Table 1 shows the impurity levels for various Pfizer antibodies using the processes outlined in Figure 1. The impurity profiles are collected after the final chromatography step. The percent monomer is determined by size exclusion high performance liquid chromatography, using two G3000 SWXL columns (Tosoh, Montgomeryville, PA) in series with detection of the antibody at 214 nm. HCP is detected using a high-sensitivity sandwich Enzyme-Linked ImmunoSorbent Assay (ELISA) kit specific for either CHO or NS0 proteins following the manufacturer's instructions (Cygnus Technologies, Southport, NC). We measured residual recombinant protein A levels using a sandwich ELISA kit from Repligen (Waltham, MA). Finally, residual total DNA was evaluated by extraction using the DNA extractor kit from Wako (Richmond, VA), followed by detection of the single-stranded DNA using a threshold kit (Molecular Devices, Sunnyvale, CA).

The results indicate that for the representative antibodies shown for the different process permutations, the current purification scheme as outlined is sufficient to produce an acceptable product. A few of the antibodies required minor changes to the platform process to produce the desired product purity, as explained in Table 1. Figure 3 shows the overall virus reduction achieved for the early-phase representative antibodies in the different platform processes. Again, the results indicate that all processes achieved satisfactory viral clearance. However, the current process is advantageous as we increase column capacities to accommodate increasing titers, as insurance for acceptable virus clearance results.

Table 1. Impurity profiles and classifications for various therapeutic monoclonal antibodies using the platform processes outlined in this article 

The current process, as shown in Figure 1, provides Pfizer with a combination of purification and virus clearance robustness, allowing us to quickly move products into the pilot plant for clinical manufacturing, thus accelerating progress toward the clinic. Even though the other processes shown in Figure 1 also met these criteria, they did not meet our new standard for robustness established by increasing product titers. A process was needed that was more adept at handling changes in the composition of the cell culture broth as well as unexpected increases in titer following scale-up, requiring additional column capacities. Although the current process bucks the trend of reducing the number of unit operations, it increases the chance of meeting the purification needs of a greater number of projects in a shorter time frame. These early-phase purification processes can be optimized further once proof of concept has been achieved. 

As cell culture titers continue to increase, the biopharmaceuticals industry will be faced with new challenges, including greater product heterogeneity and increasing impurity levels. As scale increases for early-phase manufacturing, resin capacity must increase to minimize operating costs, and  therefore, it will be necessary to carry out studies to determine the impact of these changes on virus clearance and the removal of impurities. If increased capacity cannot meet these needs, alternative separation methods such as simulated moving bed chromatography will become paramount. 

It also will become necessary to increase the throughput of nanofiltration devices, so that they do not become the next process bottleneck.  In addition, we must develop new technologies for ultrafiltration to meet the need for high-concentration drugs used for subcutaneous injection. These are just some of the challenges facing the industry today. 

The antibody purification platform process used in Pfizer's Global Biologics organization has undergone changes over the years in response to increases in both production scale and cell culture titers. These changes have resulted in a purification scheme that takes advantage of previous knowledge to reduce development time, while ensuring that product purity and virus clearance are sufficient for clinical manufacture. The goal is to manufacture early-phase material without having to perform downstream development, and the current process as outlined in Figure 1 is capable of meeting that goal. However, further increases in titer will likely bring more challenges, and thus it will be necessary to implement additional modifications to our process to keep pace with increasing upstream productivity. 

We would like to acknowledge the entire purification group in Global Biologics for their help, as well as the cell culture groups for their provision of the cell culture broth necessary to complete these studies. In addition, these studies would not have been possible without the support of the process development analytics group.

 is an associate research fellow, all at Pfizer Global Research and Development Global Biologics, Bioprocess R&D, Chesterfield, MO, 636.247.6519, 

1. Fahrner RL, Knudsen HL, Basey CD, Galan W, Feuerhelm D, Vanderlaan M, Blank GS. Biotech and Gen Eng Reviews, 18 (2001), 301–327.

2. Shukla AA, Hubbard B, Tressel T, Guhan S, Low D. J Chrom B. 2007;848:28–39.

3. Low D, O'Leary R, Pujar NS. J Chrom B. 2007;848:48–63.

4. Jagschies G, Grönberg A, Björkman T, Lacki K, Johansson HJ.  BioPharm Int. 2006;Jun (suppl):10–19.

5. Glynn J, Zaman F, Chen B, Wealand J, Mensah P. Development of a MAb harvest protocol. Biochemical Engineering XV: Engineering Biology from Biomolecules to Complex Systems. Quebec City, Canada. 2007;Jul 15–19.

A purification scheme to maximize the efficiency of the purification process and product purity while minimizing the development time for early-phase therapeutic antibodies.

The Development and Application of a Monoclonal Antibody Purification Platform

Cation exchange capture chromatography is the basis of the well-established non-affinity purification processes for many human monoclonal antibodies (HuMAbs). In all these processes, a concentration and diafiltration tangential flow filtration (TFF) step is essential to adjust the processing conditions for efficient capture on the cation exchange resin. Attempts to replace this preconditioning step with the precipitation of either host cell proteins (HCPs) or the antibody itself from clarified CHO cell culture broth have been successful. The application of a host cell protein exclusion strategy on the first column resulted in a significant reduction of process contaminants while maintaining high binding capacity. This allowed robust scale-up of the precipitation-based non-affinity processes. Because the demand for contaminant removal on the post-precipitation steps of the process was minimal, the antibody was purified to therapeutic quality in just two ion exchange steps. This improvement reduces the costs of antibody purification from high-titer cell culture batches still further by addressing critical constraint points during downstream processing.

Purification processes for antibodies have improved progressively over the last 20 years so that a limited number of unit operations can now handle a more than 50-fold increase in cell culture productivity. Most current processes for therapeutic antibodies use Protein A for capture chromatography with one or more subsequent polishing steps. These affinity processes must be modified to handle increasing antibody titers. In particular, the precipitation of cell line–derived contaminants through the addition of chemicals such as caprylic acid can generate cleaner feed streams that require fewer downstream process steps.

Processes that do not incorporate affinity columns for either the capture or polishing steps have also been developed successfully. These processes generally include concentration and diafiltration steps for primary recovery so that the feed stream is conditioned for efficient capture by ion exchange chromatography.

  A TFF step used for this purpose also achieves the partial clearance of process-derived DNA. There are several alternatives to replace the primary recovery TFF step, such as expanded bed chromatography

  or precipitation of the antibody from the cell culture broth.

  Selective precipitation of antibodies can be achieved by adding polyethylene glycol (PEG) for research-scale preparations, but more recently, PEG precipitation followed by two ion exchange polishing steps was recommended for commercial-scale antibody purification processes.

  The antibody precipitation step must be scalable to ensure consistent feed quality and efficient contaminant clearance in the rest of the process.

Here we describe the successful integration of a precipitation step into non-affinity purification processes by optimizing differential binding and elution conditions for cation exchange (CEX) chromatography resins, an approach that is referred to here as the "HCP exclusion strategy."

  The comparative performance of non-affinity processes initiated with either filtration or precipitation is discussed in the context of antibody production from CHO cells at a titer of 5 g/L.

Process 1: Ion exchange process scheme with primary recovery TFF

Simple non-affinity purification processes incorporating concentration and diafiltration steps for primary recovery have been developed for many HuMAbs, and have been scaled up to work with ~100-L columns that process up to 5,000 L of cell culture broth.

 These processes include one or two ion exchange or mixed mode chromatography steps for polishing. The process begins with buffer-exchanged clarified cell culture supernatant from the TFF step, which is conditioned to allow antibody capture on the CEX resin. The resin can also clear most of the HCP and DNA by applying the HCP exclusion conditions. In addition, the high binding capacity of CEX resins (50–120 mg/mL of resin) can reduce the number of capture cycles. In the case of two-step non-affinity processes, an anion exchange (AEX) membrane is the only necessary polishing step. This step clears residual DNA, endotoxins, and HCP, and provides an orthogonal viral removal mechanism. The loading capacity of an AEX membrane is influenced by the purity of the process intermediate from the capture step, and is limited to a few g/mL of membrane because of the need for HCP removal (Table 1). However, in some of the three-step non-affinity processes, the Q membrane load can be increased to 20 g/mL of membrane without compromising its ability to remove adventitious viruses.

Process 1 can handle a large-scale 5 g/L cell culture harvest because of the efficient capture step, the low number of capture cycles, and the rapid processing time made possible by disposable membrane chromatography. However, the productivity of this purification process can be improved further by precipitating contaminants or the antibody as the initial step before capture chromatography.

Process 2: Ion exchange process scheme with selective contaminant precipitation

Protein precipitation is often applied in the food, blood-product, and enzyme manufacturing industries

  and is accomplished by the addition of salts or organic solvents, or by pH titration. Antibody manufacturers are becoming more interested in these practices because they are suitable for processing bulk products at high throughput. Precipitation technologies have the potential to reduce or eliminate the bottlenecks that will inevitably be encountered when processing antibodies at the ton scale.

  They may also lead to the development of cost-effective downstream processes that require fewer unit operations for purification.

In the current case study, a small molecule (additive 1) was added to the clarified CHO cell culture supernatant and the pH was adjusted to precipitate most of the HCPs. The resulting precipitate was removed by depth filtration, leaving behind residual HCP in the feed. Scaling up the precipitation step is subject to variations in process parameters such as mixing speed, time, temperature, and pH, the results of which are better controlled when CEX conditions are optimized for differential binding, washing, and elution of the antibody. Transition from the precipitation step to CEX chromatography by applying the HCP exclusion strategy enables the effective integration of two-step non-affinity process schemes. Using this protocol, HCP levels can fall below 10 ng/mg after the first column (Figures 1 and 2). This degree of purity cannot be achieved in a single step when using Process 1, with diafiltration as the primary recovery step. Product quality even exceeds that achieved with Protein A as the capture step. Precipitation, combined with low pH conditioning, can prepare the feed stream for CEX chromatography with a binding capacity of up to 100 mg/mL resin, minimizing the number of process cycles while simultaneously providing virus inactivation. The material from capture is then processed by AEX membrane chromatography to remove negatively charged contaminants and adventitious viruses. Because the purity of antibodies eluting from the first column is so high, tens of grams of protein can be loaded per mL of membrane, making the AEX step faster and more economical for the application of disposable unit operations (Table 1). 

Process 3. Ion exchange process scheme with selective antibody precipitation

Antibodies can be precipitated by the addition of salt or polyethylene glycol, and the precipitate can be separated from the bulk feed by centrifugation or filtration. In the current study, the antibody was precipitated with additive 2 and the precipitate was dissolved in a resuspension buffer with a pH and conductivity suitable for direct loading onto a CEX column. Based on the known behavior of HCPs on CEX resin, we were able to design a clearance strategy that reduced the level of HCP to 40 ng/mg of antibody. When further processed by AEX membrane chromatography, HCP levels fell below the detection limit, making a two-step non-affinity process feasible (Figure 3). Mixing, incubation time, and the temperature during precipitation must be controlled to reduce variation in the load material and to achieve consistent recovery and quality. Resuspension of the antibody precipitate requires approximately 30% of the original clarified bulk volume, therefore significantly reducing the processing time for multiple CEX cycles. The protein load on the AEX membrane is determined to ensure the robustness of the process at large scale. Dilution of the protein from the CEX column is sufficient for loading on the AEX membrane chromatography, which can be operated at very high flow rates. 

Comparative analysis of the three processes

In Process 1, a relatively stable in-process intermediate is produced after the diafiltration step with a log reduction value (LRV) of ~3 for DNA relative to the clarified cell culture supernatant. For some antibodies, the capture step reduces HCP levels to below 100 ng/mg, allowing the development of a two-step non-affinity purification scheme. For others, HCP levels remain above 500 ng/mg after CEX and the process may require more than one polishing step. Therefore, to convert the majority of processes to two-step non-affinity schemes, either direct contaminant precipitation or selective antibody precipitation is necessary. Because the additives used for precipitation are small molecules that are generally accepted in protein purification processes, additional clearance steps are not necessary as the additives are removed in the first column's flow-through and washes.  

In process 2, depth filtration can separate the bulk feed containing the target protein from the precipitated contaminants, and this bulk can be loaded directly onto the first capture column. There is no reduction in the harvest volume from the precipitation step compared to TFF, so the CEX cycle time is extended during loading. Fast-flow high-binding ion exchange resins can compensate for this increase in process time, however, especially for low-volume, high-titer processes (>25 g/L) where further dilution may not have a significant impact.

  Precipitation does not require any specialized equipment and is easily transferable to existing large-scale manufacturing facilities. Further purification of the antibody by CEX chromatography achieves consistent product recovery and quality (Figure 2). With reference to HCP removal after the first column, process 2 was more efficient than processes 1 and 3 for many HuMAbs. Process 2 also takes the best cost advantage of disposable chromatography.  

In process 3, the direct precipitation of the antibody by additive 2 (based on its hydrophobicity) also results in significant contaminant removal. However, this technique is prone to greater variation during scale-up and HCP levels are higher in CEX-purified material. Although the contaminant level was high, a single Q membrane polishing step was sufficient to reduce HCP to undetectable levels. The qualitative nature of the contaminants differs substantially between process 3 and process 1. The amount loaded onto the Q membrane is much lower than in process 2 (Table 1) to accommodate the scalability and robustness of the two-step non-affinity process. Overall, the process exploited hydrophobicity as well as CEX and AEX orthogonal separations to produce therapeutic quality material. The advantages of process 3 include volume reduction, faster processing time, and easy adaptation to facilities equipped with centrifugation or gravity filtration.

Table 1. Productivity comparison of the three non-affinity purification schemes 

Protein precipitation is an efficient replacement step for primary recovery TFF and can be integrated easily into the two-step non-affinity purification process for many antibodies. The precipitation-based process can help minimize the cost of antibody therapy by increasing productivity and reducing operating costs. Similar viral clearance strategies can be implemented for both ion exchange processes with either diafiltration or precipitation of contaminants (Table 2). Process 2 has the potential to clear adventitious agents by a LRV of >4 during the precipitation step.

  Significant virus removal was accomplished by membrane chromatography in addition to other orthogonal viral inactivation and removal methods to produce therapeutic-grade antibodies.

Table 2. Viral clearance comparison for three non-affinity purification schemes 

A scalable and robust non-affinity process scheme with two ion exchange process steps provides a further opportunity to reduce the manufacturing costs for recombinant antibodies and thus the cost of goods. The two processing steps can be combined into an entirely membrane-based process scheme. The successful operation of contiguous anion and cation exchange membranes was recently reported for a commercial antibody manufacturing process.

  Even in the case of the slightly lower binding capacity of CEX membranes, the high flow rates can still offer further economic and operational advantages.

 is senior director of process development, 

 is assistant director, purification process development, and 

 is a senior process engineer, purification process development, all at Medarex Inc, Bloomsbury, NJ, 908.479.2451, 

1. Glynn J. Process-scale precipitation of impurities in mammalian cell culture broth. BioPharm Int. 2008 Mar (supplement):12–18. 

2. Glynn J. Development of a precipitation alternative and improvements for Protein A for impurity reduction in clarified broth containing a monoclonal antibody. BioProcess International Conference; 2008 Sept. 23–26; Anaheim, CA. 

3. Wang J, Diehl T, Watkins-Fischl M, Perkins D, Aguiar D, Arunakumari, A. Optimizing the primary recovery step in non-affinity purification schemes for HuMabs. BioPharm Int. 2008 Mar (supplement): 4–10.

4. Noel R, Lihme A, Lim J, Mol O, Sinclair A. Disposable adsorption systems for monoclonal antibody processing: practical, scalable operations for multi-product facilities. Recovery of Biological Products, 2008 June 22–27; Quebec City, Canada. 

5. Ramanan S, Stenson R. Method of isolating antibodies by precipitation. International patent WO/2008/100578. A2. 2008 Aug 21.  

6. Arunakumari A. Extending the efficiency of cation exchange bioseparations by novel approach for high capacity Humabs manufacturing. Recovery of Biological Products; 2008 June 22–27; Quebec City, Canada.

7. Arunakumari A, Wang J. Purification of human monoclonal antibodies: non-Protein A strategies. In: Gottschalk U, editor. Process Scale Purification of Antibodies. Hoboken, NJ: John Wiley & Sons, Inc.; 2009; p. 103–123.

8. Arunakumari A, Wang J, Ferreira G. Alternatives to Protein A: Improved downstream process design for human monoclonal antibody production. Biopharm Int. 2007 Feb (supplement): 36–40.

 9. Ingham KC. Precipitation of proteins with polyethylene glycol. Methods Enzymol. 1990;182:301–6. 

10. Gottschalk U. (2006) The renaissance of protein purification. BioPharm Int. 2006 June (supplement): 8–9. 

11. Low D, O'Leary R, Pujar NS. Future of antibody purification. J Chromatogr B. 2007;848:48–63. 

12. Percivia press release: DSM and Crucell announce record achievement in PER.C6 Technology. Heerlen/Leiden, the Netherlands; 2008 June 16. 

13. Seng RL, Lundblad JL. Viral inactivation process. United States patent US 4939176. 1990 Dec 20. 

14. Giovannoni L, Ventani M, Gottschalk U. Antibody purification using membrane adsorbers. BioPharm Int. 2008 Dec;21(12):48–52. 

In three non-affinity purification processes based on cation exchange capture with high binding capacity, applying a host cell protein exclusion strategy enabled robust scale up and better economics.

Advances in Non-Protein A Purification Processes for Human Monoclonal Antibodies

The question of whether a plant is capable of processing a given titer is, of course, an oversimplification. It is the rate of mass flow into the process that determines the ability of a downstream facility to purify the output of the reactors successfully. In this article, an example plant with fixed equipment was used to suggest where the limitations to plant throughput occur and how better process design and new methods have enabled improvements to be made over the last five years. The article also considers which technologies are, and in the future will, be available to push these limits still further.

It would be difficult to attend a conference or read an article on the development or production of monoclonal antibodies (MAbs) without being made aware of the predicted bottlenecks in the downstream processes handling the high-titer fermentations now being achieved. Facilities designed to purify the output of fermenters of a given size become unable to cope with the mass of product required of them. Where possible, increased column sizes and membrane areas can be used, but at the largest operational scales physical constraints and the prohibitively high costs of retrofitting mean that reducing the number of batches is the only realistic, though undesirable, option. 

It may not be necessary to produce antibodies at such large volumes because the same amount of product mass can be obtained from a smaller plant, but from a historical perspective similar arguments have been used for the production of antibiotics and to satisfy the global requirement for Protein A in processes for the purification of MAbs.

  Therefore, this article will assume that there is, or will be, the need for high-titer, large-volume fermentations to supply the market for MAbs. 

Models are useful tools to define process performance and gain insight into process fit in the confines of an existing plant. They are, of course, only an approximation—a model of the real situation limited by the information available for input. Models are as varied as their outputs, and are often constructed to answer a question or range of questions, which explains some of the variability seen in the conclusions reached in the literature.

The goal of this model was to understand how changing titers affected a largely idealized production plant in its ability to match the purification output with that of the fermenters. This means that the numbers given in terms of fermenter titer represent the maximum that can be processed in the plant before the downstream operation limits the number of batches that can be produced per annum. 

If we consider a plant constructed to produce MAbs for in-market supply and apply limitations to its operation, we can start to see where the bottlenecks occur, and begin to identify techniques and technologies to target the causes. In this case, the major limitations of the model are described in Table 1. 

Table 1.  Major assumptions of the model 

Table 2 summarizes the evolution of the process, taking us from how processes were developed around five years ago, through today's modern processes, and how these might change in the future using some of the techniques that are beginning to be adopted by the industry. 

Table 2.  Explanation of process evolution 

For an in-market supply production plant, the use of disposables is more limited than at smaller scales of operation. The financial incentives and physical limitations for using plastic over steel are often high, particularly for chromatography and tangential flow filtration (TFF) operations. There are still areas in the plant where disposables can be used effectively as long as the space and flexibility has been pre-engineered into the facility. 

In this instance, the use of disposables is limited to adding buffer hold vessels to the new process. After the processing time bottlenecks have been overcome, the supply of buffer becomes limiting. For this plant, freeing up a large buffer hold vessel by supplementing the hold tanks with 2,000 L of disposable storage enables batches of up to 4.3 g/L to be produced without any bottlenecks (Table 3). 

Table 3.  Describes where the first bottleneck occurs in each step of the process evolution 

This technique, developed by Pall Life Sciences (East Hills, NY) and based on patented technology, can be thought of as TFF in a direct flow mode.

  One of the main applications of this technology is its use as a fully disposable retention ultrafiltration system for early-phase material manufacture in flexible, low-capital-cost plants. Lonza (Basel, Switzerland) wanted to test the technology for application in large-scale manufacture for the in-line concentration of significant volumes of process streams to allow them to fit in the confines of existing intermediate product hold tanks. The premise of the system is shown in Figure 1—flat sheet membranes are manifolded together to create a serpentine flow path of progressively smaller membrane areas. This maintains the cross-flow across the membranes while the feed stream volume (and hence volumetric flow rate) is reduced. 

Figure 1.  A diagrammatic representation of a stack arranged in single pass mode. The stages are separated by special silicone gaskets, which are displayed here in dark grey to add clarity. 3 x 2 means 3 cassettes in parallel, 2 cassettes in series. 

New techniques can greatly improve the MAb purification process.

BioPharm International-03-02-2009

Economic Drivers and Trade-Offs in Antibody Purification Processes

Purification Strategies to Process 5 g/L Titers of Monoclonal Antibodies

The Development and Application of a Monoclonal Antibody Purification Platform

Advances in Non-Protein A Purification Processes for Human Monoclonal Antibodies

A Purification Platform for the Production of MAbs from Fermenters with Titers of 5 g/L and Beyond

Platform Technology for Developing Purification Processes