It is common practice to freeze protein bulks for long-term storage, allowing product life to be extended up to several years. For high-volume products such as antibodies, the batch size can be hundreds of liters. Freezing such large quantities in a timely manner has generally been handled using "freeze–thaw vessels", which are jacketed metal tanks with a cooling surface provided by internal coil and fin assemblies. With the increasing cost of high-grade steel alloys, it is desirable to explore disposable technology alternatives to costly freeze–thaw vessels. The following is a report on the design and testing of a prototype large-scale bag freezing system developed at Genentech. 

Interest in disposable technologies for long-term frozen bulk storage has been on the rise. This is partly because of the high capital cost of large freeze–thaw vessels, the scarcity of higher alloys as a material of construction, differences in pressure vessel fabrication standards, and requirements from country to country. Disposable containers are typically available in presterilized and custom fitted configuration, alleviating the need for clean-in-place and steam-in-place steps required for traditional freeze–thaw vessels.  

Polymeric containers with capacity of up to 20 L can be frozen in conventional freezers, though many days may be required before the entire volume reaches the desired temperature. The Celsius, a commercial system offered by Integrated Biosystem Inc. (a division of Sartorius Stedim Biotech, Goettigen, Germany), allows rapid freezing (several hours) of material in flexible containers at up to 16.5 L per container with the ability to process up to six containers at once. However, the system requires specialized equipment in addition to the freeze–thaw skid and is not operationally optimal for large batch sizes. 

The lack of a commercial large-scale disposable container system that could meet our needs prompted us to develop the technology in-house. The following reports our efforts at designing and testing the first prototype of a large scale freeze–thaw system based on disposable bags that would complement our current frozen bulk storage strategy. We discuss our rationale and design methodology, and present experimental results from the first at-scale prototype. 

Disposable bags used as liquid bulk storage containers are readily available in volumes of 1,000 L and above. The technology is proven and, because of its high structural flexibility, is relatively resilient against mild impact from blunt objects. Unfortunately, these soft-wall containers are much less forgiving when the contents are frozen. Light bumps may compromise the integrity of the bag, and the puncture may not be discovered until the contents are thawed. Such difficulties must be taken into account when designing a freezable bag system. Our aim was to develop a system that is compatible with facilities already geared toward handling commercially available freeze–thaw tanks. We had four main design constraints: 

Each individual container must hold at least 100 L. There is potential for large bulk sizes, possibly in excess of 1,000 L per batch, and the use of many small containers would cause increased handling, tracking, and testing. 

The system must be compatible with current commercially available freeze–thaw skids. The system also must be able to function under the same conditions with operational times, equivalent (within a few hours) to those of current commercial freeze–thaw vessels.  

The system must be compatible with the bulk transportation and storage infrastructure used by commercial freeze–thaw vessels. The footprint and vertical dimensions must not be larger than those of existing 300-L freeze tanks in order to accommodate storage freezers, air shipping containers, and entrances to existing facilities. 

The system must cost less than current commercial freeze–thaw tanks. 

A large-volume freestanding or minimally enclosed bag would be difficult to handle and offer insufficient protection against puncture. A solution to this problem is to use the bag as a liner inside a non-disposable container. This "bag in a can" approach offers puncture protection as well as blockage against vapor transmission though the bag wall. The basic incarnation of this type of container would be a simple, thin-walled metal shell which would be placed in a freezer and rely solely on convected cold air for freezing the contents. Based on computer simulations, however, the freezing time for such a device would be unacceptably long. A better solution would be to fully jacket and insulate the container, similar to a traditional freeze tank, and use the current freeze–thaw skids to provide the require cooling.  

The cost of the jacketed bag holder would be significantly higher than a simple enclosure. This configuration is necessary to achieve a sufficient heat transfer rate to freeze the bag contents in an acceptable amount of time. An additional benefit of this design is that the jacketing assembly provides thermal insulation protection during transport. Furthermore, because the product does not contact the metal surface, the container or shell can be fabricated from a low grade, less costly, metal alloy.  

Table 1.  Characteristics of bag shapes depicted in Figure 1. 

Various container shapes and dimensions were considered during the initial design phase of the project. Figure 1 depicts several such containers. The rationale behind our selection or dismissal of each candidate is summarized in Table 1. We conducted transient simulations of the freezing process for water in the basic shapes shown in Figure 1 using fluent computational fluid dynamic software with the solidification model. Figure 2 shows the results of the simulations in the form of remaining liquid fraction as a function of freezing time for each of the basic shapes. Also plotted for performance comparison purpose is the Fluent simulation result for the commercially available Integrated Biosystem Inc., (IBI) 300 L CryoVessel. In that case, the freezing time obtained experimentally was within 15% of the simulation value. These simulations represent the best possible cases because we did not account for resistance caused by the bag material or the contact resistance stemming from microscopically imperfect contact between the walls of the container and the bag.  

The results from the simulations show that for these basic shapes, the overall freezing performance follows the cooled surface area to volume ratio, as expected. The IBI 300 L vessel has a wetted surface area to volume ratio of 0.157 cm

  but this value includes contributions from the internal fins, which are not active cooling surfaces. Although composite shapes, as shown in Figure 1e, provide more cooling surfaces, the increased complexity in manufacturing both the holder and the bags negate the benefits of these configurations.  

To satisfy the design requirements, we selected a rectangular shape (Figure 1d) with dimensions that would accommodate a 100-L fill volume (the actual volume is larger to account for expansion of the ice). In the proposed system, three bags can fit into a three-cavity jacketed holder that provides the same capacity and occupies the same footprint as the IBI 300 L CryoVessel. Figure 3 shows the dimensions that were specified for the manufacture of the first prototype holder and bags. The bottom angle of five degrees allows for drainage of the bag contents. Each vertical wall was purposely designed with a one-degree outward tilt from bottom to top to ensure that the bag contents freeze from the bottom up. The dimensions were chosen with consideration to manufacturability and performance. Compromises had to be made to satisfy all constraints. For example, narrower cavities would yield faster completion of freezing but would be more difficult to fabricate.  

The prototype bag holder (Figure 4) was fabricated in accordance with the dimensions presented in Figure 3. Each cavity is independently jacketed and connected in parallel to a common cooling loop inlet and outlet. A drain opening is present at the lowest point in each compartment to permit fluid removal during cleaning. The final production unit should have a cover for each cavity that would provide additional moisture blockage as well as anchor points for the hoses and accessories from each bag. Additionally, a single large insulated lid would cover the top of the bag holder for thermal protection.  

Matching bags with a nominal capacity of 117 L each were manufactured by Thermo Fisher Scientific (Logan, UT), in the 3-D configuration from HyQ CX3-9 film. 

Preliminary performance testing was conducted using an IBI CU5000 freeze–thaw skid. The freezing set point was –50 °C, and the thawing set point was 25 °C for 12 hours, then stepped down and held at 5 °C until stopped. 

The bags were inserted into the cavities and gently inflated with compressed air to expand all the folds in the plastic film before filling the bag with liquid. The first set of experiments was conducted with deionized water at the 100-L fill level. It was discovered that the jacket surrounding each cavity did not extend sufficiently close to the top, resulting in long freeze and thaw times for the top 2–3 cm layer of liquid. This problem can be resolved in a subsequent prototype holder by designing deeper cavities to account for the gap in the jacketing. For the current prototype, all subsequent experiments were conducted with a 90-L fill volume. 

The prototype bag provided a port fitted with a flexible weighted internal hose intended as the dip tube for fluid introduction and removal. A similar arrangement without the weight was to serve as the thermowell (in this case, the hose end was welded shut) that can accept a probe for temperature monitoring during the freezing and thawing operation.  

After the bags were filled with liquid, we discovered that the flexible thermowell tended to coil up and did not provide adequate placement accuracy, which could severely affect temperature monitoring. Furthermore, a flexible thermowell would not allow for the removal or re-insertion of the temperature probe after the bag contents have been frozen. To a lesser extent, a similar problem affected the dip tube. In that case, the dip tube failed to reach to the bottom of the bag, leaving a considerable amount of residual liquid after emptying of the contents. Consequently, we decided to use rigid polycarbonate (PC) tubes for the dip tube and thermowell. A custom split bar assembly was used as a clamping mechanism to hold the PC tubes vertically. The split bars were secured to the metal bag holder by adhesive-backed industrial Velcro, which allowed minor adjustments in the position and easy removal when necessary. Figure 5 shows the setup with the rigid dip tube and thermowell. Recirculation mixing can be accomplished by pumping the liquid from the bottom of the bag by the dip tube and returning it near the top at the opposite side of the cavity, approximately 10–12 cm above the liquid level. The outlet for the return port is aimed toward the nearest wall to minimize potential foaming.  

The tops of the bags were removed and the sides were secured to the cavity walls using heavy-duty adhesive tape to monitor the progress of freezing in each cavity. A video camera connected to a computer was used to record images at one-minute intervals as a time-lapse movie during the freeze and thaw phases of each experiment. This allowed us to visually assess the completion of the freeze or thaw process accurately. 

We conducted a series of freeze–thaw experiments in the prototype bag holder filled with water as well as a 25 mM citrate buffer containing 150 mM sodium chloride (NaCl). Sodium chloride solutions form a eutectic at approximately –21 °C; as a result, the NaCl solution exhibited very different freezing dynamics compared with pure water. Table 2 summarizes the results for a select number of runs, which are discussed in a subsequent section. The completion time for freezing or thawing was determined from the time-lapsed video data. In some instances, it was not possible to clearly observe all three cavities simultaneously; in those cases, no time data is given. Freeze time is defined as the time required to freeze all of the liquid; thaw time is the time required to melt all of the ice as visually determined from the time lapse movies. The freeze–thaw skid permits the recording of only one external temperature probe. The location of the probe varies from experiment to experiment and is indicated in Table 2.  

A flow rate of 1 L/minute was used in all cases involving recirculation mixing during thaw. Similar conditions are adequate for post thaw mixing in freeze–thaw vessels. Only the cavity containing the temperature probe was connected to the pump. It should be noted that the recirculation time of three hours was chosen arbitrarily; the optimal duration will be determined later but is expected to be between three and five hours. 

Experiment D, in which no bags were used, established that the heat transfer performance of all three cavities is reasonably comparable. The freezing of 3 x 90 L of water was completed in eight to nine hours (for this specific geometry, the simulated freeze time estimated using Fluent software is about nine hours), although the right-hand cavity was ~10% slower than the other two. For comparison, an IBI 300 L CryoVessel can completely freeze 300 L of water in 10–11 hours on the same freeze–thaw skid. Although in this experiment the bag holder is not operated in the intended configuration, this can be viewed as being the worst case scenario in terms of heat load on the freeze–thaw skid because there is no additional thermal resistance caused by the bag material or contact resistance. The results suggest that the heat transfer fluid flow is distributed fairly uniformly among the three cavities, although the right-hand cavity may be slightly more restricted.  

A new set of bags was installed immediately preceding experiment A1. All of the remaining experiments, listed chronologically in Table 2, used the same set of bags. The heavy-duty adhesive tape used to anchor the top of each bag to the holder walls also provided some protection against moisture caused by condensation seeping between the bag and the holder walls. A thin water film present between the bag and cavity wall can significantly enhance the heat transfer rate by minimizing contact resistance. This can be seen by comparing the freeze time data for the center cavity in duplicate experiments A1 and A2. A closer inspection of the center cavity confirmed that large wall areas had been wetted. A comparison of the freeze time between the left and center cavities in experiment A1 also reveals that the performance varies from cavity to cavity, with the left cavity performing significantly worse than the center cavity (more than 16 hours versus 13.5 hours). In this instance, the discrepancy can be attributed to the poorer fit of the left side bag (no bag wall contact in some areas) compared to the center bag. The bags used in this study were manufactured by a manual process; hence, the dimensional tolerances are rather wide at ± 25 mm. 

In experiment set B, the PC thermowells and dip tubes were added to each cavity. This addition did not have an effect on freezing performance. We also investigated the magnitude of heat transfer enhancement from having a wetted bag–cavity wall interface by intentionally introducing water in this area. In experiment B2, we injected water between the bag and the cavity wall of the center cavity. In this case, there was no noticeable decrease in freezing time as compared with B1, probably because that interface was already wetted from the condensation generated in previous freeze–thaw cycles. Injecting water in the interfacial region between the bag and wall of the left cavity, however, resulted in significant improvement in freezing time, as seen in experiment B3. The magnitude of this improvement is consistent with those seen for the center cavity in experiment set A. The thin layer of water reduced the contact resistance between the bag and cavity wall.  

The temperature traces for experiment set B are shown in Figure 6 and give a measure of reproducibility from experiment to experiment. Note that the freeze or thaw time for the right -hand cavity does not vary appreciably, indicating that there was minimal moisture ingress between the bag and cavity wall throughout the studies. Hence, the right cavity serves as an internal control for the various experiments.  

In experiment sets B and C, the cavity monitored by the temperature probe also underwent recirculation mixing during the last three hours of thaw. Interestingly, the thaw times show little variation across all experiment sets, ranging from approximately nine to 10.5 hours, irrespective of cavity location, contact resistance, or recirculation. In IBI 300 L vessels, 300 L of water can be thawed in approximately eight to nine hours using the same freeze–thaw skid. Unlike the freezing process, controlled by conductive heat transfer where the thermal resistance increases as ice is formed, the thaw process is primarily dominated by natural convective heat transfer. As thaw proceeds, the heat transfer rate between the container walls and the contents does not change drastically. 

In experiment set C, the contents of the center cavity were replaced with 90 L of buffer. The freezing time was greater than that of pure water by two to three hours (compared to experiments B and C, center cavity). This reflects the lower thermal conductivity of the solute containing ice and the freezing point depression from the cryoconcentrate. Similar results are observed for standard freeze–thaw vessels. Experiment C2 also included thermowell sleeves, constructed from the same material as the bags. A configuration involving thermowell sleeves integral to the bag instead of the original flexible tubing would allow the use of a rigid thermowell made from any convenient material without introducing another product contacting surface. This is shown in Figure 5. There were no significant differences in the measured temperature resulting from the addition of thermowell sleeves (data not shown). 

Figure 7 shows a sequence of images taken during freezing and thawing from experiment C1. It shows that during thaw, a piece of ice is kept at the bottom of the cavity by the thermowell but eventually fractures and floats to the top. The ice motion is unpredictable and ice in close proximity to the heated wall will melt faster, as expected. This is likely a major contributor to the variability in thaw times.  

In Table 3, we have summarized several important technical issues that were identified during the testing of the prototype large freeze bag holder and offer some possible solutions.  

Articles

A solution for the problems of a "bag in a can" system would be a fully jacketed and insulated container, similar to a traditional freeze tank.

Design and Testing of a Prototype Large-Scale Bag Freeze–Thaw System

The closed and controlled environment of single-use technologies in a biopharmaceutical production process can significantly reduce processing time. A fully disposable option eliminates cross contamination, cleaning, and subsequent validation, reducing the use of water for injection. Lower operating costs, smaller equipment footprints, increased productivity, and rapid turnaround time are some of the major benefits that are driving the market demand for single-use technology in manufacturing. This article discusses a contract manufacturing organization's perspective on the use of disposable technology in the biopharmaceutical production process.

To eliminate contamination and to comply with the FDA cleaning regulations,  manufacturers spend a lot of money, time, and resources on equipment cleaning and sterilization—steps that require large volumes of water for injection (WFI)—and cleaning validation. Most bioprocess equipment is made of stainless steel and requires assembly, clean in place (CIP), and steam-in-place (SIP), all of which result in manufacturing downtime. By using disposables, contract manufacturing organizations (CMOs) can save WFI costs by not having to clean the stainless steel bioreactors and validate the cleaning process every time.  

These reasons have fostered the advent of single-use technology, which has been embraced by technologically inclined  companies. These companies understand that significant investment capital is needed to construct a manufacturing facility. It costs $10 million to construct, equip, and launch a 100-L traditional pilot plant facility. The cost goes up to $40 million for a 1,000-L pilot plant and for a 10,000-to 20,000-L facility, the cost becomes prohibitive, and may be as high as several hundred million dollars.

Because of their customizable design, disposables facilitate faster design, construction, and commissioning of facilities and offer manufacturing flexibility—a significant operational benefit.  Reduced processing time, enhanced productivity, smaller and flexible production facility, and reduced operating costs are some of the major benefits that are driving the market demand for single-use technology. 

Contract biomanufacturing companies are continually striving to improve and optimize the process of manufacturing biotherapeutics at every stage, starting from preclinical toxicology material to large-scale current good manufacturing practice (cGMP) commercial production. A contract manufacturer on  average makes a 33% profit in manufacturing a biotherapeutic.

  In order to be competitive and be commercially viable, CMOs have to make improvements in this area. They have a pressing need to economize the manufacturing process, skillfully manage their project pipeline, and help their sponsors accelerate their speed to clinic or market.  

Process economics is not for just the dedicated CMOs, however. A biopharmaceutical company with inbuilt capacity is in a similar situation, spending more than 20–25% of its operational cost on manufacturing,

  and is equally interested in lowering the production costs. Disposable technology presents an attractive opportunity to cut down production costs for both biopharmaceutical and contract manufacturers. 

Where Are Disposables Used in a Biomanufacturing Process? 

Single-use filters, single-use tangential-flow filtration (TFF) membranes, flexible tubing, sterile liquid containment bags, disposable bioreactors, mixing systems for media preparation, sterile bags with normal-flow filter manifolds, and aseptic connectors are all used in appropriate steps during the process. These disposables optimize processes and accelerate biomanufacturing production time. In downstream processing, disposable formats are used for cell removal, harvest clarification, membrane absorption chromatography steps, viral filtration, ultra filtration, and diafiltration. At Laureate, we use single-use technology at all of these steps throughout the biomanufacturing process.  

For small-scale culture scale-up, we routinely use disposable flasks and roller bottles. For larger scales up to 1,000 L, we use disposable Wave cell bags. 

Wave disposable systems range from 1 L to 500 L in capacity, and we use them in conjunction with disposable cell bag 20/50, 100/200, or 500/1000 Wave bioreactor systems. These are customized systems that use rocking motion to mimic the agitation produced by stand-alone systems. They have built-in feedback controls to monitor carbon dioxide, dissolved oxygen, pH, and other parameters. Our studies have shown that cell growth in these systems for most cell lines (Chinese hamster ovary [CHO] cells, BHK, hybridoma) is comparable to that obtained from stand-alone systems (data not shown). We observed similar growth trends throughout the run when we scaled up from a 20-L to a 1000-L wave bag  (Figure 1). 

Another type of disposable bioreactor uses hollow-fiber (HF) cartridges. In such perfusion bioreactors, cells grow inside cartridges, around a semipermeable 10-kD hollow fiber membrane. Cells are fed and waste is removed by perfusion across the membrane. HF disposable bioreactors facilitate a plug-and-play approach to cell culture, and have the advantage of concentrating the product produced by the cells.  

Various vendors supply bioprocess containers (BPC) for solution preparation. These containers are made of polyethylene, ethylene vinyl alcohol (EVOH), and other films. They are available in a variety of sizes ranging from 25 mL up to 500 L, and we transport them in re-usable barrels or totes. BPC systems are completely configurable for various uses, including direct connection to chromatography systems or process tanks. The bioprocess containers are tested to United States Pharmacopeia (USP) Class 6 standards before use. The vendor performs the following tests and confirms validation at the time of purchase:   

cytotoxicity (agar diffusion and elution) 

Disposables are an integral part of membrane chromatography for protein purification. We use cation and anion exchange filters from various vendors to replace  cation and anion exchange resins and columns. The cost analysis shows that disposable membrane filters are substantially more cost-effective than stand-alone columns and resins. In addition, by using these filters we have eliminated the need for packing and testing columns as well as long-term storage of materials.  

Anion exchange filters are typically flow-through (FT) systems that bind rDNA, endotoxin, residual protein A, and host cell proteins, and we use them for polishing steps. A comparison study of Q filters from two different vendors (Figure 2) shows that disposable membrane filters generally result in >90% yield postprocessing. We have observed similar yields when we run process-scale columns packed with Q anion exchange resins. 

Cation exchange filters act as product-binding filters similar to cation exchange resins and bind IgG and Factor VIII. Cation exchange filters, however, have lower capacity than most resins and are not efficient for intermediate purification steps. 

We use different types of disposable filters mentioned below for virus removal (Table 1). 

Table 1.  A comparison of four types of filters commonly used in a post-chromatography purification process for virus removal. Flux rates, recovery, and convenience of use were studied for each filter for different processes.  

In a disposable HF filter, the product is ideally filtered through the hollow fiber membrane by pressure at 14 psi. Total yields are greater than 95% and membranes are scalable to 4.0 m

 . However, a pressure tank is necessary if pressure feeds are used and therefore, this system is not fully disposable in its ideal configuration. We have observed that this system can function at 120–150 L/H/m

  and can achieve greater than 6.7 log removal of polio virus (vendor study). 

TFF membrane filter cartridges are composed of three disposable membrane modules connected in parallel. However, stainless steel membrane holders are needed for setup and therefore, this system is not fully disposable as it has nondisposable parts. The TFF system functions at 50–80 L/H/m

  and achieves greater than 6.0 log removal of retrovirus (vendor study).  

Polyvinylidene difluoride (PVDF) filters are completely disposable cartridge filters that can be hooked up directly to pumps or chromatography systems. The product is pumped through the filter in a single or parallel channel configuration. The filters are scalable to 30 inches and can be connected in tandem to polishing columns, if needed. We have observed that these filters function at flow rates ranging from 5–20 L/H/m

 . The PVDF filters are commonly used in the industry and have successfully removed greater than 6 logs of retrovirus in our processes and in other commercial applications. However, backpressure issues are common, resulting in slower and less efficient virus removal steps.  

Polyether sulfone (PES) cartridge filters are also available for viral removal steps. PES filters are hooked up to pumps or chromatography systems in the same fashion as PVDF. Scalable to 30", these filters are completely disposable and can be hooked in tandem to polishing columns, if applicable. Our studies confirm that PES filter flow rates are higher than PVDF filters at 90–180 L/H/m

 . We performed a virus validation study using PES filters by pumping virus-spiked product through small-scale cartridge membranes at >30 L/m

  for all runs. Results show that the product spiked separately with either 1% or 5% (v/v) of four viruses (Reo, MVM, MuLV, or PRV) behaved in a manner consistent with our scaled-down processes. In all cases, the flow-rate curves were linear, suggesting that the filters never exceeded the virus binding capacities. Studies also confirmed greater than 6 log reduction in all of the above named viruses.   

For aseptic filling of liquid parenteral products we use completely disposable sterile bags as bulk product containers and have seen excellent recovery. One caveat to using disposable bags for aseptic filling is that the users must confirm that the product solution is fully compatible with the plastic material of the bag. Our linear peristaltic pump systems have completely disposable parts, including hoses and filling needles to accommodate vials from 0.2 to 100 mL in size with fill volumes of 0.1 to 100 mL (Figure 3). 

One of the challenges of adopting single-use technology is that not all cell lines are compatible with disposable bioreactors.

Single-Use Technologies—A Contract Biomanufacturer's Perspective

Disposables are increasingly being used in the manufacture of biopharmaceuticals. This article describes the design of a fully disposable process for the cGMP manufacture of clinical trial grade plasmid DNA. It addresses the rationale for implementing such a process with respect to the manufacture of patient-specific plasmid DNA vaccines for the treatment of leukemia. The process incorporates a number of disposable technologies, which are simple to use and thus reduce the need for investment in expensive equipment and cleaning validation. 

Disposable technologies are being used either as replacements for conventional technologies or for the development of new processes for the manufacture of current good manufacturing practices (cGMP) grade biopharmaceuticals. These technologies are particularly applicable to manufacturers of investigational medical products (IMPs) where the range of products required to be manufactured is often diverse and at a small scale. In this article, we discuss the design of a disposable platform for the manufacture of patient-specific plasmid DNA (pDNA) vaccines for the treatment of leukemia. 

The potential for treating a range of diseases by the delivery of DNA in vivo is being realized in the clinic in several different areas such as cancer, infectious diseases, and autoimmune diseases. The advantages of using naked pDNA over viral delivery routes include low immunogenicity (allowing a course of multiple treatments) and a simpler manufacturing system.

  The pDNA vector cassette is highly suitable for the rapid development of various vaccine targets, such as personalized vaccines. One limitation for pDNA vaccination is the relative efficiency of the targeting and delivery of pDNA, which is being currently addressed by improving delivery modes by either chemical or physical means (e.g., adjuvants, liposomes, gene gun, or electroporation).  

The Clinical Biotechnology Centre (a part of the UK's National Blood Service) has manufactured a range of pDNA vaccines for academic collaborators, and those vaccines are currently undergoing Phase 1 and 2 trials. Since 2005, we have been involved in the manufacture of anti-idiotypic bespoke pDNA vaccines for patients suffering from multiple myeloma. This B-cell malignancy is characterized by the secretion of a serum immunoglobulin paraprotein, which can be identified molecularly and re-engineered into a single-chain Fv construct. The construct is cloned into the pDNA vector cassette containing an immunostimulatory tetanus toxoid fragment.

  Vaccination with the pDNA encoding the patient's paraprotein should induce the patient's immune system to mount anti-idiotypic responses against the tumor. The pDNA construct from one patient to the next may only differ by less than 2% of the backbone, making it suitable for designing a manufacturing platform. The vaccination regimen requires less than 50 mg of pDNA to be manufactured. 

The promise of being able to deliver personally tailored vaccines for patients is a big challenge for the biopharmaceutical industry. Often, investments in such "orphan technologies" are not necessarily favored over more conventional generic approaches, where cGMP issues are less complex and the expected returns are greater. An ideal platform process would address the need to reduce manufacturing costs and allow rapid turnaround of batches of product.  

Processing costs can be significantly reduced by eliminating the capital outlay in expensive, dedicated equipment (such as fermenters or chromatography systems) and opt for a process that is both disposable and simple to perform by an operator. A single-use process also eliminates the need for equipment decontamination procedures and cleaning validation studies, which are cumbersome and costly.  

Some prerequisites are necessary to underpin the platform process to ensure compliance with current regulatory requirements. The appropriate sourcing of key materials is critical, such as the exclusion of animal-derived ingredients and the use of pharmaceutical-grade (USP, class VI) disposable plastics throughout. The prudent selection of the appropriate E. coli host cell and pDNA vector will also maximize plasmid quality and yields.

The use of validated and pharmacopoeia-compliant quality control assays is essential to demonstrate the effectiveness of the process to purify pDNA away from process-related impurities.

  The platform should be validated according to the cGMP guidelines and achieve a high quality of product to satisfy the current regulatory requirements.

An outline of the design of the cGMP process for manufacturing pDNA vaccines is shown in Figure 1. Each step should be attributed a specific function. 

Plasmid DNA is transfected into E.coli by heat-shock, and clones containing the pDNA are identified under a selective pressure present on the vector (such as an antibiotic marker or by regulation of an essential E. coli protein). In an industrial setting, a master cell bank (MCB) is laid down from a clone, which is then cultured in large-scale fermenters by fed-batch, high-cell density culture.

  We have demonstrated that it is possible to omit the clone selection or MCB steps and culture a randomly selected clone in low-tech disposable shaker flasks without compromising high pDNA yields (>3 mg pDNA per gram of wet weight E. coli). We are able to routinely produce up to 200 mg of pDNA in a culture volume of less than 4 L in 24 hours. High specific yields are favored over high volumetric yields (mg pDNA per liter of ferment) because they reduce the relative starting load of E. coli-sourced contaminants. The ferment is harvested by centrifugation into single-use plastic centrifuge bottles at late log for maximal pDNA yields. Because of cost, we decided not to use disposable fermenter systems or crossflow applications for this step. 

Standard alkaline lysis methods must be optimized to ensure the efficient lysis of cells and subsequent removal of major contaminants such as genomic DNA and proteins.

  The lysis is performed in a plastic container containing a low-level outlet attached to prefiltration and 0.22 μM filtration devices. The whole assembly and container are autoclaved together before use. Addition of a secondary salt such as ammonium acetate or calcium chloride precipitates and removes RNA (up to 45% reduction in contaminating nucleic acid load observed).

  The secondary salt partitions the flocculent (precipitated material) in an upper layer away from the lysate, which enables the subsequent clarification of the lysate by dead-end filtration (Figure 2). The clarified lysate can be stored in a bioprocess container or can be pumped directly into the crossflow filtration container for the start of the purification step. 

Figure 3 shows the complex nature of the lysate, which is known to contain large amounts of RNA, endotoxin, and trace amounts of contaminating genomic DNA, protein, and other non-supercoiled pDNA related isoforms. The pDNA constitutes less than 5% of the total mixture, which presents a challenge to purify the desired monomeric supercoiled pDNA to homogeneity from contaminants that have similar physiochemical properties. Supercoiled pDNA is tightly coiled and is widely accepted as being the therapeutically efficacious form, mainly due to its relatively small size compared to other isoforms.

The purification step initially involves the diafiltration and concentration of the lysate with a hollow-fiber crossflow filtration cartridge. This device is assembled with a vacuum-resistant plastic container fitted with a series of three-way valves, vent filters, and Pharmed tubing in an enclosed system. The buffers, which are stored in bioprocess containers (BPCs), are attached to a three-way valve on the diafiltration line (Figure 4). An appropriate molecular weight cut-off membrane is chosen (300 kDa or smaller) to allow the retention and high recovery of supercoiled pDNA from the lysate. The SC nature of plasmid requires the selection of a membrane with a MW cut-off approximately 10 times smaller than expected to prevent losses across the membrane. Carefully defined operating conditions are also necessary to reduce the effect of shear, because plasmid is prone to affects that are exacerbated by the prevailing high salt conditions. The lysate is diafiltered, concentrated, and filtered into a BPC before anion exchange chromatography. Crossflow filtration of the lysate has been previously reported to be an excellent RNA removal step with more than 90% clearance demonstrated in practice (Figure 3).

Anion exchange chromatography media in packed columns is the conventional approach for the initial capture of pDNA. The negatively charged pDNA molecules bind electrostatically to the media, but limitations in the binding capacity is a problem that remains unresolved, mainly because of the large size of the plasmid molecule. This has recently been addressed by a shift in emphasis from beaded media to a membrane approach. Membrane devices offer many benefits, including disposability (no need for column packing and qualification), an increased dynamic binding capacity for larger molecules, and high process flow rates.

  In our platform, a Mustang Q (Pall Corporation, East Hills, NY) capsule is used downstream from the crossflowed lysate (in contrast to other organizations that have used this step as a direct capture from E. coli lysate).

  Pretreatment by crossflow ultrafiltration is advantageous for increasing the Mustang Q's binding capacity for pDNA (since the RNA no longer competes once removed), improving recovery of eluted pDNA, and achieving higher product purities. The membrane device is attached to Pharmed tubing and three-way valves for processing during purification (Figure 5). The apparatus is autoclaved, operated in a Grade A laminar flow hood by a pump, and all fluid movements are contained in BPCs. If the step has been validated in terms of the wash conditions, elution volumes, and flow rates, the cartridge does not need to be attached to expensive instrumentation. The loading and washing of the sample is rapid, followed by a longer and staggered multiple high salt elution step. This ensures that the recoveries of pDNA achieve greater than 70% of total plasmid load, because membranes are characterized by slower off rates for pDNA.

  Mass balance analysis across this step efficiently removes 2 log

 orders of endotoxin and more than 50% of the remaining residual protein. 

The eluate is collected into a BPC and ammonium sulphate is added to the BPC to achieve a final concentration of 1–1.5 M for the negative capture hydrophobic interaction step. At a set concentration of salt, impurities such as endotoxin (>90% removal) and RNA (>95% RNA removal) are selectively bound onto the HIC media while pDNA is recovered. The media is preconditioned with ammonium sulphate, autoclaved, and pumped into the BPC to achieve a specific nucleic acid ratio load to resin. The batch-mode HIC is incubated on a rotating mixer before 0.22 μM filtration into another BPC to remove the resin and recover the product.  

Purified pDNA is formulated into the buffer of choice by a disposable scaled-down version of the previous hollow fiber membrane assembly (Figure 6). The pDNA is formulated at the desired concentration in the appropriate solution (normally isotonic) requested by the sponsor. The choice of filter membrane is also critical to avoid unnecessary losses of plasmid by nonspecific binding. 

The final product is fully tested, and satisfies current recommended guidelines (Table 1). Limited in-process QC testing is performed because of the continuous nature of the process. 

Table 1.  Final QC release testing and specifications for clinical grade pDNA 

The continuous process is suitable for the rapid turnaround required for the preparation of these vaccines at a low cost. It is aseptic, contained, and completely disposable to avoid product cross contamination. It has been estimated that on current demand, with two operators in two cleanrooms, up to 40 individual vaccines could be manufactured annually. 

By careful design, it is possible to eliminate some of the main obstacles and limitations normally associated with conventional processes (e.g., to reduce the reliance on high-tech equipment and to eliminate the use of column chromatography). The process has been shown to achieve a recovery of pDNA of greater than 45% from bacterial lysate with the quality of the final product satisfying current regulatory requirements. 

 is a business development manager, all at the Clinical Biotechnology Centre, Bristol, UK, 011.792.89388, 

 1. Schleef M, et al. DNA Pharmaceuticals. Wiley-VCH Verlag GmBH & Co KGaA, Weinheim;2005. 

 2. Stevenson FK, et al. DNA vaccines to attack cancer. PNAS. 2004;10(2):14646–14652. 

 3. Werner RG, et al. pDNA—from process science to commercial manufacture. Pharm Technol Eur. 2002;March;34–40. 

 4. European Pharmacopoeia Supplement 5.6. Gene transfer of medicinal products;2007. 

 5. Rules and guidance for pharmaceutical manufacturers and distributors. Pharmaceutical Press, London;2007. 

 6. Lee SY. High cell density culture of Escherichia coli. TIBTECH. 1996;14:94–105. 

 7. Birnboim HC, Doly JA. A rapid alkaline extraction procedure for screening recombinant plasmid DNA. 1979;7:1513–1523. 

Disposables are increasingly being used in the manufacture of biopharmaceuticals. This article describes the design of a fully disposable process for the cGMP manufacture of clinical trial grade plasmid DNA. It addresses the rationale for implementing such a process with respect to the manufacture of patient-specific plasmid DNA vaccines for the treatment of leukemia. The process incorporates a number of disposable technologies, which are simple to use and thus reduce the need for investment in expensive equipment and cleaning validation.

Disposable Process for cGMP Manufacture of Plasmid DNA

A team assessed the means to improve the production process of a polyclonal antibody from mouse serum at the 200-L production scale. The objectives of the study were to improve the current process by reducing processing time, increasing yield and purity, and decreasing operation costs. An integrated disposable assembly was built, incorporating disposable technologies for:

1. Clarification and bioburden reduction.

2. Affinity purification (using a prepacked column).

4. Final filtration (using an in-line filter connected to a bag).

The solution reduced labor costs, accelerated turnaround process time, and ensured high purity. This study was a successful first step; the proposed next step is an assessment of changes to be implemented at process scale, specifically to implement tangential flow filtration (TFF) for concentration, and to replace the final dialysis step with diafiltration.

The increasing demands on productivity in biologics processing have led companies to seek out process efficiency more broadly instead of considering only yield. Manufacturers now seek to improve batch and campaign changeover time to allow more flexible multiproduct operations. This change in views has led to increased demand for disposable technologies, which significantly reduce the changeover time between batches or campaigns. This increased demand for disposable technologies applies to the manufacture of biologicals used in diagnostics and biotherapeutics. In this study, we evaluated disposable technologies throughout a mouse gamma globulin (MGG) process to improve overall productivity in the manufacture of this diagnostic. 

The current method used to produce MGG is a batch approach that is described below. For clarification and bioburden reduction, glass fiber–poly-propylene capsules (1–1.2 μm) were used in prefiltration. Intermediate filtration was conducted using polypropylene–cellulose acetate–polyethersulfone capsules (0.65–0.45 μm). Bioburden reduction was performed using cellulose acetate–polyethersulfone capsules (0.2 μm). 

Chromatography media (ProSep-vA Ultra, 5 L) at a dynamic binding capacity of 20 mg/mL was used for the chromatography step. Two hundred liters of media were processed in a batch-wise manner with a target step yield of 90 g and 95% purity. The following buffers were used for the bind, wash, and elute steps:  

1. Binding buffer: PBS 0.01 M, NaCl 0.15 M, azide 0.1%, pH 7.1 

2. Wash buffer: PBS 0.01 M, NaCl 0.15 M, azide 0.1%, pH 7.1  

3. Elution buffer: glycine HCL 0.2 M, azide 0.1%, pH 2.8 

4. Neutralization buffer: tris 0.5 M, NaCl 0.5 M, azide 0.1%, pH 10.2 

5. Cleaning: acetic acid 2 M, ethanol 20%, after 300 L. 

For concentration and dialysis, a stirred cell concentrator with a 30 kD molecular weight cut-off (MWCO) PM membrane or a 20 kD MWCO cellulose acetate membrane were used to concentrate purified MGG to at least 35–60 mg/mL. Dialysis was performed using tubing (10–14 kD MWCO) at 2.8 °C with PBS 0.1 M, NaCl 1.5 M, and azide 0.1%, pH 7.1 for at least 24 hours with three changes of dialysis buffer. Final filtration was performed using a double membrane filter (0.8/0.2 μm). Final filtration though a 0.2-μm filter was conducted on the same day as the fill-and-finish operation. The target concentration was 25–50 mg/mL with a minimum purity of 95%. 

Described here is an experimental approach to improve the purification of MGG from serum at a 200-L production scale. The objective was to purify mouse MGG with an integrated disposable assembly at a 3% scale that balanced availability of feedstock, and to provide an appropriate scale of operation to accurately assess the scaled-down operation. 

The integrated disposable assembly (Figure 1) was built using neoprene tubing (Pharmed, Miami, FL) of suitable lengths and with the appropriate connectors attached. These were placed in autoclave bags and sterilized at 121 °C for 20 minutes. The filters were aseptically connected to tubing assemblies in a laminar flow hood. 

Chromatography buffers were sterile filtered in bags using 0.2 μm filters (Opticap XL3) with Millipore Express SHF membrane in the following quantities: equilibration buffer: 50 L; elution buffer: 10 L; and regeneration buffer: 50 L. The bags were aseptically connected to the assembly as shown in Figure 1. 

A 32 x 250 mm column (Millipore Vantage, Billerica, MA) was manually packed offline with chromatography media (ProSep-vA Ultra, 200 mL) in 20% benzyl alcohol. Packing linear flow velocities were 1,000 cm/hr. Before applying the feedstock, the column was washed with 10 CV of elution buffer, followed by 10 CV of binding buffer. The column was then aseptically connected to the disposable assembly as shown in Figure 1. 

The TFF system was assembled, and the 30 kD membrane (Pellicon 3, 0.11 m

 ) was rinsed with 10 L of sterile water (Milli-Q) to remove any preservatives. The system was sanitized with 5 L of 0.5 M NaOH. After sanitization, the assembly was rinsed with sterile water until a neutral effluent pH was measured from the cross flow and permeate lines. The assembly was then aseptically connected to the eluate bag as shown in Figure 1. 

Frozen mouse serum (6 L) was thawed then pumped into the disposable assembly. The flow rate was slowly ramped up to an inlet pressure of 10 psi. All of the serum was allowed to pass first through the clarification filter (Opticap XL5 capsule with Polysep II 1.0/0.2 μm cartridge), followed by the sterile filter (Millipak 200 gamma gold 0.22 μm filter) into a 10-L bag. The clarification filter was changed twice because of by-plugging. 

The serum was processed in a continuous operation in the assembly. A volume of 800 mL was pumped onto the column (ProSep) at a flow rate of 60 mL/minute. The void volume for the tubing and column was found to be approximately 100 mL. The column was washed with equilibration buffer (10 CV) at a flow rate of 120 mL/minute. The bound MGG was eluted with 3 CV of elution buffer at a flow rate of 60 mL/minute. The pH of the eluate was neutralized by aseptic introduction of the neutralization buffer. The column was regenerated with 10 CV of regeneration buffer and re-equilibrated with 10 CV of equilibration buffer. Regeneration and re-equilibration were both done at a flow rate of 120 mL/minute. These steps were repeated eight times to process the entire 6-L batch. The total volume used for neutralization of 6 L of eluate was 15 mL. 

The purified MGG collected in the eluate bag was pumped into the TFF assembly. The polyclonal MGG was concentrated 16X and constant volume diafiltered with PBS 0.01 M, NaCl 1.5 M, pH 7.4. 

After concentration and diafiltration, the purified MGG was pumped across a sterile filter (Opticap phillic filter with Durapore, 0.22-μm membrane) into a bag. A sample was drawn from the final purified MGG to determine the concentration and assess the bioburden of the purified MGG. The purity of the polyclonal MGG was evaluated by running polyacrylamide gel electrophoresis, followed by estimation of purity using QuantiScan software (Figure 2). The bioburden of the purified MGG was determined by plating a sample on growth media and checking for microbial growth. The yield was calculated based on 3% scale results. 

Figure 2 shows the purification profile of the polyclonal MGG from mouse serum using the integrated disposable assembly. Comparison of the integrated assembly purified MGG (well 10) to the control reveals that the purity in well 10 was very high. One band each of the light chain and the heavy chain of the MGG were observed and contaminant bands seen in the control were absent. Purity percentages were determined using QuantiScan Software. 

The proposed change from concentration/dialysis to ultrafiltration/diafiltration resulted in a significantly faster process and reduced process time from two weeks to 25 hours thus improving turnaround times and productivity at the facility. 

The changes to chromatographic buffer composition for both the equilibration buffer (from 10 mM PBS, pH 7.2, to 20 mM PBS, pH 7.4) and elution buffer (from glycine HCL, 0.2 M, pH 2.8, to glycine HCL, 0.1 M, pH 2.8) had the following benefits: an increase in final concentration to 79.28 mg/mL (minimum expected target concentration 25–50 mg/mL); 100% purity (minimum expected target 95%); and an increase in yield of 96.64% (minimum expected target 90%). Reducing filter hold up losses by adopting system rinse and air blow down may increase yields further.  

The elimination of the azide also has potential for refinement caused by contamination issues with the dilution buffer. Future work needs to be carried out in order to resolve the contamination issue in the dilution buffer. Using the purification process described, the overall cost of operation for a 200-L batch was estimated to be $41.13/g of MGG. 

 is a consultant for business and knowledge development, 

 is a senior manager for manufacturing operations, 

 is a director for strategy and corporate development, all at Millipore Corporation, Billerica, MA, 781.533.2233, 

Comparison of the integrated assembly purified MGG to the control revealed that the purity if the MGG was very high.

Antibody Purification with an Integrated Disposable Assembly

The implementation of single-use systems—which can deliver significant value through added flexibility, improved production yields, and cost savings—is gaining momentum in the biopharmaceutical manufacturing industry. The initial single-use systems that gained market acceptance consisted of tubing, bags, filters, and connectors. Today, additional types of disposable technologies are being used throughout the production cycle. This article discusses how single-use transfer lines can be incorporated into seed train and final fill applications as well as suite-to-suite transfer operations.

Manufacturers across industries rely on innovative technology to meet two critical production requirements, speed and cost-efficiency. With the increasing pressure to reduce time to market and costs, biopharmaceutical manufacturers are exploring new technologies to meet these challenges. 

With the costs to manufacture a single drug approaching $1 billion and time-to-market ranging from eight to 12 years, biopharmaceutical manufacturers are under tremendous pressure to commercialize new drugs faster at a lower cost.  

To meet these intense market demands, manufacturers are being forced to continuously boost manufacturing speed and efficiency by relying on innovative technology that can be easily integrated into their existing production processes. One solution that continues to gain momentum is single-use systems, which can deliver significant value through added flexibility, improved production yields, and significant cost savings. 

The initial single-use systems that gained market acceptance consisted of tubing, bags, filters, and connectors. These systems were used in bioprocessing facilities for process storage applications and sterile cell culture media. Single-use bioreactors then entered the market in research and development laboratories and rapidly moved into pilot plants and larger-scale production facilities as integral systems for seed train scale-up and production.   

Now, single-use transfer lines are emerging as yet another option for manufacturers to save time and cost. Unlike hard piping, the flexible tubing incorporated into single-use transfer lines allows manufacturers to quickly change process steps or convert to a new product without costly and time-consuming cleaning and validation. This is a key advantage for multiple-product facilities in which process requirements change depending on the drug being produced. Innovative manufacturers now incorporate single-use transfer lines in seed train and final fill applications as well as suite-to-suite transfer operations. 

The key benefit of single-use transfer lines is the ability to boost productivity and accelerate the time to market by reducing the downtime associated with equipment cleaning and validation. Between each production batch, fixed tubing and re-usable valves must be cleaned to maintain desired sterility. Single-use systems are presterilized and eliminate the need for traditional cleaning and sterilization. This reduced downtime translates into greater productivity and throughput. Instead of the entire process being placed on hold for validation, a single-use system can allow the process to be up and running sooner, thereby increasing output and accelerating the time to market.   

Additional cost savings result from reduced labor, chemical, water, and energy demands associated with cleaning and validation. Not only do single-use systems reduce costs, they also improve the safety of drug development and delivery. Presterilized, single-use assemblies reduce the risk of cross contamination that may lead to product loss or reduced yields. This benefit is further magnified for companies that produce multiple products within single facilities. 

Modern bioprocessing facilities feature production bioreactors with capacities of 5,000, 10,000 and even 25,000 liters. Scaling up inoculum from a few million cells in several milliliters of culture to these production volumes is a challenge that requires aseptic transfer at each point along the seed train. Traditional bioprocessing facilities accomplish scale-up using a dedicated series of stainless steel bioreactors linked together with valves and rigid tubing (Figure 1). 

To prevent contamination between production runs, a clean-in-place (CIP) system is designed into each bioreactor, vessel, and piping line to remove any residual materials. Sterility assurance at the start of each culture is provided by a steam-in-place (SIP) system consisting of steam pipes, temperature sensors, and condensate collection piping. These CIP and SIP systems require extensive validation testing and the valves and piping contained in these systems can create additional validation challenges. In addition, CIP and SIP systems must be revalidated following significant maintenance or changes to the connecting piping and valves network. 

Recent advances in single-use system technology allow bioprocess engineers to replace most storage vessels and fixed piping networks with single-use storage systems and transfer lines. Single-use systems reduce maintenance and capital expense by eliminating expensive vessels, valves, and sanitary piping assemblies. Figure 2 depicts a 10,000-L production suite, which relies on stainless steel bioreactors, but integrates single-use technology for cell culture media storage and key transfer lines. 

Single-use media storage systems are routinely manufactured for volumes ranging from 20 to 2,500 liters. Media storage systems arrive at the bioprocess facility sterilized by gamma irradiation and often are fitted with integrated filters, sampling systems, and connectors. Using an SIP connector allows operators to make sterile connections between these presterilized single-use systems and stainless steel bioreactors for aseptic transfer of media.  

Similarly, single-use transfer lines may be used to transfer inoculum between bioreactors using either a peristaltic pump or head-space pressure for flow rates up to 30 liters per minute. Such transfer lines can reduce the number of reusable valves required for transfer, and eliminate problem areas for CIP and SIP validation. Terminating each presterilized transfer line with a single-use SIP connector provides sterility assurance equal to that of traditional fixed piping at lower capital costs. 

With the acceptance of single-use bioreactors, some bioprocess engineers are using these systems for both seed trains and small-scale production. Single-use bioreactors range in volume from 1 to 1,000 liters and use two modes of mixing: rocking platform and impeller. Rocking platform bioreactors consist of pillow-style bags that are secured to a tray mounted on pneumatic lifters. Impeller-based single-use bioreactors are based on three-dimensional drum bags fitted with a single-use impeller before sterilization. These systems are connected to a cell culture media storage bag (either by aseptic welding or aseptic connectors) using flexible tubing.  

Similarly, flexible tubing with aseptic connections are used as transfer lines from one reactor to the next. For production volumes over 1,000 L, a single-use bioreactor must ultimately seed one or more stainless steel bioreactors. SIP connectors incorporated into the single-use bioreactor design can link the single-use and stainless section of a seed train. Figure 3 shows a model facility that incorporates single-use media storage and bioreactors, using stainless steel bioreactors for the two largest vessels of 2,000 L and 10,000 L. 

After upstream production, the medium containing protein must be aseptically transferred to a different location in the production facility for the final filling operation. This suite could be in a room next door or much farther away. Traditional bioprocessing facilities accomplish this transfer by using stainless steel with piping or reusable hoses as transfer lines. This equipment requires validated CIP and washing procedures before use and sterilization before each media transfer.  

Process engineers are now incorporating single-use transfer lines between the bioreactor in the process suite and the transfer vessel to simplify this process. By using presterilized connectors and tubing, the medium can be moved from the production suite to the preparation suite without the need to sterilize stainless steel piping or equipment.  

In addition to faster production, these single-use systems also give the manufacturer greater flexibility to determine which process to run in production suites. The enhanced mobility eliminates many of the restrictions that hard-plumbed piping can place on the manufacturing facility. 

The risk of cross-contamination in the suite-to-suite transfer process is high, especially in multiproduct facilities.  Presterilized, single-use assemblies reduce these risks and improve the speed and safety of drug development and delivery. Such assemblies are used in processes requiring a flow rate of less than 30 liters per minute, but more recently, suppliers have increased the diameter of single-use tubing and SIP connectors to enable bioprocessing facilities to incorporate single-use transfer lines into larger and larger production processes. 

The final step in the production process is transferring the product from the transfer vessel or bags into smaller vials, bottles, or containers for distribution. In the past, the final fill operation consisted of stainless steel equipment connected by reusable valves, rigid tubing, and steel pipes. This equipment also requires validation and must be subjected to a CIP cycle after each filling cycle is completed. Today, many process engineers are designing this operation with single-use transfer lines to reduce sterilization time and cost.  

One example of integrating single-use systems in a final fill operation is to simplify mobile stainless steel transfer tanks. Final fill tanks are designed to transfer formulated product from formulation suites to storage areas and ultimately to filling suites. To allow sterile connection to and from these vessels, designers add three-way valve assemblies to fill and drain ports to facilitate SIP operations. The design of these three-way valves makes it difficult to validate cleaning procedures. In addition, these valves require regular maintenance and may add significant weight to mobile vessels, especially in tanks with multiple inlet and outlet ports. Replacing these three-way valve assemblies with single-use transfer lines eliminates cleaning validation and maintenance steps while reducing mobile vessel weight by tens of kilograms.  

Single-use transfer lines can be attached before vessel SIP sterilization with SIP connectors (used as either steam access or condensate drainage sites), or steamed separately, just before fluid transfer. For vessel outlet, combining a number of single-use components into the transfer line can create a very robust system to ensure product safety. For example, outlet transfer lines could incorporate a single-use SIP connector to attach to the sterile holding tank. Then, a through-the-wall fluid transfer system is used to bring a portion of the transfer line into an isolator where filling occurs. Next, a quick-connect fitting or aseptic connector is used to attach the transfer line to the filling machine.   

Figure 4 depicts a mobile transfer vessel with multiple single-use inlet lines and a complete outlet line system. 

As more manufacturers take advantage of the benefits of single-use systems, their integration with traditional stainless equipment will continue to grow. All biopharmaceutical manufacturers are able to benefit greatly from single-use systems, but biotech start-ups can gain additional operational advantages by saving time and expense in the design, building, and validation of new facilities. Manufacturers also benefit by retrofitting existing facilities with single-use transfer lines to increase scheduling flexibility, production capacity, and improve production yields with minimal expenses. 

Single-use transfer lines are not limited to upstream or downstream processes and the benefits can be seen in all operations (large or small), or new and existing facilities. Whether it's connecting within a process or across different processes, this is a technology with bottom line advantages throughout the manufacturing operation.   

 is the bioprocess business unit manager at Colder Products Company, St. Paul, MN, 651.645.0091 

Single-use systems reduce maintenance and capital expense by eliminating expensive vessels, valves, and sanitary piping assemblies.

BioPharm International-11-02-2007

Design and Testing of a Prototype Large-Scale Bag Freeze–Thaw System

Single-Use Technologies—A Contract Biomanufacturer's Perspective

Disposable Process for cGMP Manufacture of Plasmid DNA

Antibody Purification with an Integrated Disposable Assembly

A Next Step in Implementing Disposables: Transfer Lines