Articles

Issue PDF

Pharmaceutical Technology North America, Bioprocessing Supplement 2011 (PDF)

The dendritic cell (DC), the most powerful antigen-presenting cell in the immune system, has been a popular choice as a basis for personalized or autologous cellular immunotherapies. For these autologous DC immunotherapies, a batch of drug product is generated for each individual patient using his or her own cells. Scientists use three main approaches to obtain or generate DCs for a patient.  

One approach is to isolate circulating DCs directly from the blood or from white blood cells collected through leukapheresis. The number of circulating DCs in the peripheral blood is extremely low, which limits the potential yields and doses that can be obtained using this approach. The advantage of this approach is that the DCs require minimal manipulation once isolated. Scientists essentially load the DCs with the antigen or antigens of interest against which an immune response is desired.  

The two other approaches involve isolating precursor cells and culturing them to generate DCs. These precursor cells are either hematopoietic stem cells (HSCs) or monocytes. HSCs require longer culturing times to generate DCs compared with monocytes and require the patient to be mobilized (e.g., pretreatment with granulocyte colony-stimulating factor) before the leukapheresis. The advantage of the HSC approach is the ability to proliferate the cells before differentiation. Monocytes reduce culture time and manipulations compared with HSCs with the limitation that the cells have with no proliferation capacity. Therefore, the key to autologous DC processing from monocytes is managing losses and optimizing recoveries at each step to ensure an efficient process. 

Generating a dendritic-cell immunotherapy

Using monocytes, Argos Therapeutics developed a robust method for generating its autologous DC therapy from leukapheresis for clinical trials in renal-cell carcinoma (RCC) and human immunodeficiency virus (HIV) indications (see Figure 1). The monocytes are isolated from the leukapheresis using the Elutra Cell Separation System (Caridian BCT). This instrument uses elutriation, also known as counterflow centrifugation, to fractionate the cells in the leukapheresis based primarily on cell volume, thus providing a final fraction of enriched monocytes. These monocytes can be cultured immediately to generate DCs, or can be frozen to be thawed and cultured when the antigen is available. Monocytes are cultured with granulocyte-macrophage colony-stimulating factor (GM-CSF) and interleukin 4 (IL-4) for 5 days to generate immature DCs in culture bags. Maturation media containing tumor necrosis factor α, interferon γ, and prostaglandin E

 is added, and the DCs are cultured overnight to mature before the antigen is introduced.  

Figure 1:  Overview of an autologous dendritic cell-manufacturing process for oncology and infectious-disease indications.  (ALL IMAGES ARE COURTESY OF THE AUTHORS)			

Antigen is added to the DCs in the form of ribonucleic acid (RNA) using electroporation. The antigen RNAs are amplified for each patient from samples of his or her tumor or virus (1, 2). This method provides antigen that is unique to that individual and tailors the DC immunotherapy for that individual. This technique, however, makes the manufacturing more complex than alternative approaches in which the antigen is universal for all patients and can be manufactured in bulk quantities. The RNA is manufactured for each patient by isolating the total tumor or viral RNA from the patient's sample. That isolated RNA is converted to first-strand cDNA through reverse transcription and amplified by polymerase chain reaction (PCR) using nonsequence-specific primers and methods, thus making them universal for all samples. RNA is generated using the resulting amplified cDNA through an 

 transcription (IVT) reaction and is post-transcriptionally capped to ensure a high capping efficiency. Using this process, milligrams of amplified RCC messenger RNA (mRNA) are generated consistently from micrograms of total RNA. For HIV, RNAs for the quasispecies of gag, vpr, rev, and nef proteins in the viral sample are amplified (2). From isolated viral RNA, the concentration of which cannot be measured by standard spectrophotometric methods, milligrams of RNA for each antigen are amplified for each patient. 

In addition to the amplified RNA from the patient's disease, an RNA-encoding cluster of differentiation 40 ligand (CD40L) is added to the RNA payload. The purpose of adding this CD40L RNA is to provide the CD40–CD40L ligation signal required by the DC to induce IL-12 secretion (3–5). IL-12 is linked to the functionality of the DC because IL-12 secretion is one of the three signals required for a typical adaptive immune response (6). A technique for quantifying the release of IL-12 from the DC immunotherapy is in development as the drug product's potency assay (7).  

This CD40L RNA is generated in bulk from a plasmid with one batch of CD40L RNA used for several batches of the DC immunotherapeutic drug product (8). For each batch of CD40L RNA, the plasmid is linearized, and uncapped CD40L RNA is generated using IVT methods. The uncapped CD40L RNA is capped and polyadenylated to generate the final CD40L RNA that is added to the RNA payload during electroporation. This process of maturing the DCs before electroporating them and adding CD40L RNA to the RNA payload has been called the postmaturation electroporation CD40L or the PME CD40L process (9). Dendritric cells resulting from this maturation process expand the central and effector memory T cells (CD8

) associated with favorable clinical outcomes (10). 

Following electroporation with the amplified RNA from the tumor or viral sample and the CD40L RNA, the DCs are cultured for 4 h with GM-CSF and IL-4 to recover, translate the RNAs, and process and present the resulting tumor or viral peptides. After culture, the DCs are harvested, formulated in autologous plasma collected during leukapheresis and cryoprotectants (i.e., dimethyl sulfoxide and dextrose), and frozen in multiple vials. Each vial is a single dose of drug product. These vials are stored cryogenically and shipped individually to the clinical site for administration to each subject. Implementing this cellular process based on elutriation, culture bags, and the PME-CD40L maturation method, yields a mean number of doses produced per batch greater than 20 for the RCC and HIV indications (see Table I). This method provides multiple years of dosing for a patient from a single leukapheresis.  

Table I: Results of the cellular process based on elutriation, culture bags, and PME-CD40L maturation methods for manufacturing clinical-scale batches of RCC and HIV.			

The drug-product release testing includes post-thaw total viable cell count and viability to verify the dose strength and immunophenotyping for identity (see Table I and Figure 2). Cell-surface markers CD80, CD86, CD83, and CD209 identify the cells as mature DCs with the appropriate co-stimulatory molecules to generate an immunostimulatory T-cell response. CD14 is a monocyte marker; therefore, the low percentage confirms that the monocytes were converted to DCs. Human leukocyte antigen-DR indicates the presence of major histocompatibility complex Class II receptor for peptide antigen presentation. These data demonstrate the consistency in the formulation and quality of the DCs despite the significant biological variability in the starting materials. The consistency in results between batches for each disease indication and for the two different disease indications establishes that the clinical manufacturing methods are robust.  

Figure 2: Post-thaw immunophenotyping results confirmed the identity and quality of the dendritic-cell drug products generated for renal-cell carcinoma  (RCC) and human immunodeficiency virus (HIV) after implementing the cellular process based on elutriation, culture bags, and PME-CD40L maturation methods. CD is cluster of differentiation, and HLA is human leukocyte antigen.			

Automating the process for commercialization

The process for generating this autologous DC immunotherapy has been consistent and robust for the RCC and HIV indications during Phase II clinical trials. Elutriation and culture bags have moved key cellular processes to functionally closed, single-use disposable units. However, some manipulations and processing are still performed using manual methods, including open manipulations in biological safety cabinets. Although this approach may be feasible for the number of batches required for clinical trials, it is not practical for commercial manufacturing because every patient requires a new batch of material to be produced.  

Advanced RCC has a relatively low incidence of an estimated 9400 new cases in the US in 2010 (11–13). This number alone would mean a considerable number of batches to manufacture per year. An HIV product would likely require manufacturing for more than three times as many patients per year in the US, compared with an advanced RCC product. Even more staggering is the number of patients for Provenge (sipuleucel-T), the first approved autologous cellular therapy produced by Dendreon. Provenge was approved for metastatic castrate-resistant (i.e., hormone refractory) prostate cancer, an indication for which more than 100,000 patients are treated per year (11, 14). These numbers of batches per year require feasible processing methods to meet the demands of commercial manufacturing for autologous cellular therapies. 

If a company used the method described above to manufacture an autologous DC immunotherapy, which could generate years of drug product with one leukapheresis, the process scale would remain the same for commercialization. The company would not need to scale up for autologous therapies. The question would be how to scale out or address the throughput needed for commercialization. Answering this question was the goal of an automation project that developed novel manufacturing equipment. The cornerstone of the developed approach is the incorporation of single-use, functionally closed disposables throughout the process. This method was recognized as crucial for autologous cellular-therapy manufacturing to eliminate cross-contamination concerns and minimize turnaround time between processes, given the throughput needs.  

Personnel began the process by adapting the cellular processing methods to incorporate elutriation and culture bags. Equipment to perform the remaining cellular-processing steps at the scale required for autologous cellular therapies, however, did not exist. Therefore, two similar instruments were designed with the functionality to perform each of these process stages, including processing the autologous plasma collected during the leukapheresis procedure for use in drug-product formulation, and designing custom disposable sets to handle the nuances of each process. The reason for designing two instruments was to create one for plasma and monocyte processing, which are less complex. The second instrument addressed the unique needs and volumes required for mature DCs, such as the electroporation step, including handling the addition of precise volumes of RNA normalized to the DC concentrations, and formulation.  

The programming and disposables were designed so that reagents and the cellular inputs for each process were connected to the appropriate disposable set using standard functionally closed disposable manipulation methods (e.g., tube welding). The design incorporated the removal of culture bags or other process outputs using tube-sealing methods. Therefore, the cellular equipment would never be exposed to patient material, and all products and processing will be closed, thus eliminating potential contamination events and product losses resulting from open manipulations. Although it is always crucial to minimize batch failures, the need is greater for autologous cellular therapies, given that each batch is specifically manufactured for a patient, and the patient undergoes leukapheresis to provide the starting cellular material for manufacturing. Because the high variability of the biological starting material influences manufacturing success, personnel need to incorporate any possible additional controls during manufacturing to ensure the generation of product. Processing with single-use, functionally closed disposables and automated methods is a way to significantly improve process control. 

The development of the automated cellular equipment and associated disposable sets began with identifying the key processing steps and needs. Moving from elutriation to culture (or freezing to thawing and then culture) required media-exchange steps and the ability to resuspend and distribute cells in a functionally closed disposable component. Similarly, harvesting the DCs, electroporation, culturing following electroporation, and final formulation all required cell-concentration and media-exchange steps. Methods for reliably and accurately addressing this manipulation of cells were developed for even the small-volume manipulations required for electroporation or formulation (i.e., 5–30 mL) while not compromising processing time when large-volume manipulations (large volume for an autologous cellular therapy means as much as 5 L) are required. Overall, the time required for cellular processing using automated equipment is similar to that for the manual methods. The majority of the time required for cellular processing, however, is dedicated to incubation to generate DCs from monocytes, followed by culturing for maturation and culturing for recovery after electroporation.  

The biggest challenges to resolve were the management of losses and the maximization of recovery at each step. Minimizing the manipulations for harvesting procedures and media exchanges was critical to ensure that process efficiencies (which were measured by the percentage of monocytes in the initial leukapheresis resulting in RNA-electroporated, mature DCs vialed as drug product) for the automated process were similar to those of manual processing.  

Table II: Results for the four initial feasibility runs using the developed automation equipment and functionally closed disposables.			

Four initial cellular-feasibility runs performed on prototype automated equipment using the developed disposable sets demonstrated the cellular equipment's ability to perform the processing efficiently with well-controlled, small volumes (see Table II). The formulation of the drug product was on target; it had high DC viability post-thaw. The number of doses produced met expectations; the variability in dose numbers related to the variability in the number of starting monocytes present in the leukapheresis for each run. The drug-product immunophenotyping results for these automated runs confirmed identity and consistency in quality (see Figure 3). Results were similar to those generated in clinical manufacturing (see Figure 2).  

Figure 3: Post-thaw immunophenotyping results confirmed the identity and quality of the dendritic-cell products generated in the four initial feasibility runs using the developed automation equipment and functionally closed disposables. CD is cluster of differentiation, and HLA is human leukocyte antigen.			

As the cellular automated equipment and disposable sets were developed, equipment also was developed to amplify autologous RNA from a tumor sample. Though various platforms for automated equipment for nucleic acid manipulations exist, they are generally based on high-throughput methods and open manipulations of plates. For isolating and amplifying nucleic acids for an autologous therapy, this type of equipment could potentially be placed in a barrier isolator to achieve the isolation required for manufacturing. The cleaning requirements between patient samples to prevent cross-contamination, however, would be time consuming. Ensuring that existing automation platforms could withstand vaporous hydrogen-peroxide decontamination between processes would have required additional instrument development. Also, the cleaning validation would have been extremely challenging because the products generated were nucleic acids. The concept, therefore, was to use a functionally closed disposable container for processing, and to design that disposable so that patient material was never in direct contact with the equipment (see Figure 4).  

Figure 4:  Prototype equipment for automated autologous RNA processing.			

The developed RNA disposable container had two main components. The first component was a rigid tray that incorporated all that was necessary to isolate and process the nucleic acid (e.g., pipette tips, reagents, mechanism for nucleic acid isolations and purifications, and PCR plate for all incubation steps). The disposable container also includes spectrophotometer cuvettes and specially designed volumetric cuvettes required to determine the concentration and volume of the isolated and amplified nucleic acids. These cuvettes ensured that the equipment can calculate yields and the volumes required for concentration normalization. They also enabled the equipment to perform the entire process without interruption or data from an outside source.  

The second component was a flexible barrier with an incorporated pipette head. This flexible barrier was sealed onto the rigid tray to generate the closed RNA disposable. When closed, the flexible barrier enabled the six-axis robot arm incorporated in the equipment to access all areas in the rigid tray to perform the liquid transfers and other manipulations required for processing. Along with the robotic arm, the automated RNA equipment incorporated the thermal cycler needed for PCR and all incubation steps, as well as the spectrophotometer needed to determine concentration. In initial feasibility runs, the prototype automated equipment and functionally closed disposable container generated amplified RNA comparable to the amplified RNA generated using current manual clinical processing methods, thus demonstrating that this automated concept is appropriate for manufacturing drugs for oncology indications. This concept can be readily adapted to infectious-disease indications to amplify RNA from a viral sample. 

These prototypes demonstrated the feasibility of this automated approach. The automated equipment and the associated disposable sets or components provides a platform technology to enable commercial manufacturing of complex autologous DC immunotherapies. 

is a director of manufacturing and process development at Argos Therapeutics, 4233 Technology Dr., Durham, NC 27704, tel. 919.287.6300, fax 919.287.6301, 

 4. J.P. Ridge, R.F. Di, and P. Matzinger, 

 12. S.A. Karumanchi, J. Merchan, and V.P. Sukhatme, 

SEER Cancer Statistics Review, 1975–2007,

 S.F. Altekruse et al., Eds. (NCI, Bethesda, MD, 2010). 

The authors developed automated equipment that uses functionally closed disposables to perform cellular and ribonucleic processing.

Methods for the Automated Manufacturing of an Autologous Dendritic-Cell Immunotherapy

From an industrial research and development (R&D) perspective, the design and development of protein therapeutics today appears somewhat akin to the rational design of small-molecule discovery back in the 1970s when lead compounds were generated from known physiological substrates or ligands. Facing a need to find novel and diverse small-molecule leads, attention in the 1980s centered on high-throughput screening (HTS) technologies and compound libraries. Those libraries, albeit large, were hardly diverse, with most therapeutic agents coming from a few target protein classes. Complementation of libraries with natural products, the development of combinatorial chemistry, and application of focused-library sets followed. This evolution, together with automated methods for content-rich assay systems and fast make-test cycles, enhanced discovery of novel, potent, and diverse lead series.  

Contrast this with the present analagous processes for protein therapeutics: the discovery and development of novel biologics is hardly diverse, efficient or rapid. State-of-the art protein discovery and development use multiple expression hosts (e.g., mouse, 

 Chinese hamster ovary (CHO), and NS0)  and several reformatting steps between hosts are often necessary during testing, scale-up, and production. The process of developing cell-based protein expression systems that are efficient, consistent, and scalable often is difficult and sometimes impossible using currently available technology.  

To date, more than 150 protein drugs have been approved for clinical use, nearly all of which are produced in cell-based expression systems, such as 

. These cell-based systems have several limitations, and many biologics can't be developed in these systems. For example, these systems only allow the overexpression of proteins that don't affect the physiology of the host cells. For many expression systems, identifying cell lines that stably synthesize high protein titers of the desired product is a time-consuming and labor-intensive process. Ideally, the same production host for rapid variant discovery, production for animal testing, and manufacturing of a clinical candidate would be used. 

Ideally, one would want to emulate the huge leap made in iterative drug design seen in small-molecule discovery, namely, rapid make-test cycles and generation of multiple parallel libraries of drug candidates with diverse structural elements to optimize activity while maintaining feasibility for manufacture. An ideal system would do the following: 					

Make fast make-test cycles a prerequisite for re-iterative design on the order of three to five days, similar to those for focused small-molecule libraries

Create efficient and rapid expression and purification that allows for libraries of hundreds to thousands of protein-sequence variants to be simultaneously tested per make-test cycle using standard off-the-shelf robotics equipment

Incorporate preferred sequences defined from selection technologies, such as ribosome or phage display, into whole protein therapeutics for testing

Enhance the diversity of chemical structures by expanding the library of available amino acids at specifically targeted points in the protein sequence from 20 natural to many hundreds of non-natural amino-acids

Optimize several properties (e.g., agonist or antagonist, affinity, stability, and predictive manufacturability) simultaneously through rapid high-throughput make-test cycles 

Create processes that are not only rapid but amenable to rapid scale-up and cGMP manufacturing once the desired therapeutic construct has been identified.

As ambitious as such a system would seem, several exciting technologies are emerging that improve expression systems and enhance diversity to enable modification of intrinsic properties of proteins, such as enzyme catalytic efficiency or binding.  Others combine different properties in single therapeutics by conjugation chemistries. Further emerging technologies can lead to more rapid and parallel expression of many protein drug candidates. Getting all of these desirable technologies into a single amenable platform that has the flexibility to be scaled and support cGMP manufacturing is in sight. 

Early improvements in endogenous protein-based therapeutics produced new, commercially successful therapeutics with desirable properties by simply extending sequence incorporating fusion to proteins such as the constant fragment of antibodies (Fc)  or by PEGylating to increase half-life.  Beyond these early approaches, considerable effort to produce ever-more elegant constructs that combine two separate functions have been made. One promising approach, antibody drug conjugates (ADCs), involves using a targeting antibody to known tissue selective cell-surface antigens or receptors to target conjugated toxins or cytotoxic drugs and so  enhance selectivity over normal tissue.   

Successful design of effective ADCS is complex and requires linking cytotoxic drug payloads to tumor-targeting antibody constructs. The selection of an ideal antigen target for optimal internalization and specificity for tumor tissues is critical.  The design of linkers that are stable in circulation, but cleave when internalized in tumor cells to release the cytotoxic drug, adds to the complexity, but the other major technical hurdle has been to define how the cytotoxic payload with linker are conjugated to the targeting antibody.  The biopharmaceutical companies Seattle Genetics and Immunogen have developed robust platforms that depend on conjugation of linkers and cytotoxic warheads to available cysteine or lysine residues, respectively, in the tumor targeting antibody sequence.  

Despite successes with ADCs, there are many examples where seemingly optimized functional components (i.e., antigen-binding motif, linker, and drug-conjugate) do not translate into a developable therapeutic candidate. ADCs produced using conjugation chemistries to endogenous cysteines and lysines inevitably lead to the production of multiple species of the ADC with the drug conjugated in varying payloads of  between one and nine molecules per immunoglobulin G (IgG). Furthermore, all sites of conjugation are not equal. Some conjugations interfere with antigen-binding epitopes, thereby reducing binding affinity and/or drug half-life (1). All too often, poor efficacy is revealed in the clinic only after significant investment in cell-based expression systems and scale-up. 

Several technologies aim to provide chemically amenable sites within a protein sequence for the posttranslational chemical conjugation of small-molecule drugs, peptides, or other constructs to improve or add functionality. For example, sequence-specific conjugations producing homogeneous ADCs with fixed payloads are aimed at improving tumor-cell killing and increasing therapeutic index. Engineered ThioMabs (Roche/Genentech technology) that use natural cysteine residues  that must be carefully unmasked during production for subsequent site-specific conjugation have shown preclinical proof-of-concept (2). These approaches await further clinical validation.   

Carlos Barbas'  laboratory at The Scripps Research Institute exploited the use of exposed tyrosine residues within the complimentarity determining regions (CDRs) of IgG molecules as the basis for linking drug conjugates. CovX, acquired by Pfizer in 2008, was founded to develop this technology. These sorts of approaches are attractive in that posttranslational chemical coupling to a common IgG construct with resulting extended half-life represents a platform amenable to many different small-molecule or peptide agonists. 

The introduction of non-natural amino acids (nnAAs), or those amino acids not part of the 20 naturally incorporated ones into proteins, plays an important role in basic peptide and protein research. They are increasingly used to develop biologics with enhanced pharmacological properties beyond providing sites for drug conjugation. NnAAs can be introduced through chemical synthesis in peptides or biosynthetically in proteins. Currently, only peptides and very small protein drugs with nnAAs are on the market because they can be made synthetically and avoid the limitations of cell-based expression systems. A prominent example is the semisynthetic, broad-spectrum antibiotic, ampicillin, into which the nnAAs D-phenylglycine and D-4-hydroxyphenylglycine have been incorporated (3). 

The opportunities to broaden protein diversity and properties with nnAAs are enormous, as is the ability to incorporate chemical modifications in proteins that can endow current biopharmaceuticals with improved or new properties. These chemical modifications can change the characteristics of proteins, including ligand-binding properties, stability, spectroscopic properties, folding behavior, catalytic efficiency, and substrate specificity. These modifications provide possibilities to develop biobetters and biosuperiors that have superior pharmacological properties, including improved safety profiles, longer half-life, and enhanced activity (4). 

Much effort has been put into developing technologies that ensure a site-specific incorporation of the nnAAs with a high rate of yield. Various methods for site-specific introduction have been established, both semisynthetic and recombinant methods. Few methods, however, have made it from the bench at small-scale  protein production to commercial scale.  

A recently formed biotechnology company, Redwood Biosciences, is using an approach based on the work of Carolyn Bertozzi's laboratory at the University of California, Berkeley. Her work focuses on genetically encoded aldehyde tags and aims to exploit a specific sequence (originally found within the sequence of sulphatases) that is posttranslationally recognized and modified by a formyl glycine-generating enzyme to produce a so-called aldehyde chemical handle (5). The incorporation of the CxPxR sequence at specific positions in candidate protein therapeutics provides a means to produce a site-specific nnAA with a reactive aldehyde amenable to drug conjugation. 

One of the oldest methods for nnAA incorporation into proteins uses auxotrophic strains from 

that cannot synthesize a specific natural amino acid and thus have to uptake it from the growth medium. A structurally similar nnAA can be supplied within the growth medium in place of the natural amino acid and will be alternatively incorporated into the protein. A major downside is that the specific nnAA will be incorporated at every site coding for the natural amino acid and can lead to misfolding and impaired function of the target protein, or the nnAA can be incorporated in the host-cell's proteins, which can have toxic effects (6). Allozyne has pioneered this type of cell-based expression system that incorporates nnAAs into protein sequences, but this approach requires extensive re-engineering of the target protein sequence used, due to the region-specific nature of the nnAA incorporation using this method. 

Ambrx has developed cell-based nnAA incorporation systems where 

or CHO cells are engineered with orthogonal pairs of transfer (tRNA) and tRNA synthetases to charge and incorporate nnAAs at selected codons at specific points in the coding sequence of the expressed protein. This approach is a significant advance and provides answers to at least some of the questions raised about nonspecific sites of conjugations in ADCs. For truly expanding the number and variety of nnAA that can be incorporated to determine the effect on function, even at a single amino-acid position, the approach demands significant investment to engineer further orthogonal pairs of tRNA synthetases and tRNAs that can recognize a library of nnAAs. A further complication is that these pairs should be exquisitely selective over natural amino acids to avoid their incorporation over the desired nnAA, which can be challenging in a drug-manufacturing context with strict regulatory requirements. When this challenge is taken into account, along with the variability in efficiency with which nnAAs are absorbed into the cell, these systems will not likely be amenable to fast reiterative make-test cycles with libraries of nnAAs at multiple sites of incorporation.   

All of these considerations suggest a clear need to move away from the conventional cell-based protein expression systems to address the critical requirement for a rapid make-test system that is amenable to many parallel re-iterations of site-specific incorporations of defined natural amino-acid sequences or multiple nnAAs at multiple sites. The answer may not lie with cell-based systems at all, but with completely 

 biochemical protein synthesis based on novel cell-free expression systems. 

Cell-based expression systems: parallel reiterative design and scalable platforms

Adnexus Therapeutics (acquired by Bristol-Myers Squibb in 2007) has developed an 

-based platform for producing adnectins. Adnectins are derived from human fibronectin and many trillions of adnectin variants can be generated to represent a screenable library for desirable therapeutic properties. Scale-up from a selected lead is rapid, albeit with the requirement for PEGylation for clinical candidate manufacture (7). 

 Fabrus has recently addressed the rapid make-test cycle approach for biologics using arrays of predefined Fab antibody sequences produced in a high throughput expression system based on production of proteins in 

, cell-lysis, and subsequent protein purification (8). The large volumes of cell culture required for high-yield parallel production of Fabs requiring a significant investment in specialized robotics equipment. The method allows production of hundreds of variant Fab proteins over a one-week production cycle to begin variant testing in high-throughput biochemical-binding assays.  

Although cell-free protein synthesis has been practiced for decades as a research tool, only recently have advances suggested its feasibility for commercial biologics drug development and production as an alternative to conventional cell-based expression systems (9). An ideal cell-free protein production platform would produce fully soluble and correctly folded proteins at high volumetric productivities at any scale. The platform would be rapidly and predictably optimized by systems-level process design and control without the demanding requirements for maintaining cell viability and  be readily adapted to high-throughput methods, including 

evolution of proteins to allow incorporation of nnAA into polypeptides. The platform would be based on simple batch systems using standard bioreactors that are known to scale to thousands of liters for both cell fermentation and subsequent cell-free protein production (10, 11). 

Early efforts at developing such a system focused on projected costs that were much too high, as well as on proteins with disulfide bonds that could not be folded effectively. By focusing on basic biochemical reactions and controlling cell-free metabolism, these limitations have been methodically addressed (12). Amino-acid supply has been stabilized, and metabolism activated to dramatically reduce substrate costs by requiring only the addition of nucleotide monophosphates to drive energy production. Commercially available 

, wheat germ, rabbit reticulocytes, and insect-cell extracts do not offer this advantage and are suitable only for research exploration at small scale. Control of the sulfhydryl redox potential has been gained and a robust disulfide isomerase added to facilitate oxidative protein-folding (13). These advances not only suggest production feasibility for pharmaceutical proteins containing the 20 natural amino acids, but they also provide enabling technology for incorporation of nnAAs at commercial scale. 

A recent publication demonstrates that this open cell-free system (OCFS) developed by Swartz and collaborators can be optimized for high-level production of proteins to allow for scale-up to commercial levels once the target protein is identified (14). The authors expressed a multidisulfide-bonded protein, biologically active granulocyte-macrophage colony-stimulating factor (rhGM-CSF), at titers of 700 mg/L in 10 h. Importantly, they could show that the product was linearly scalable from starting materials in 96-well plates up to 100-L culture volume (14). The open nature of the system allows mass spectrometry-based profiling of the cell-free metabolome and proteome.  Rapid testing of the effects of addition and subtraction of various components for system optimization can be modeled without the requirements for tuning more complex cellular networks required for maintaining cell viability commonly encountered in mammalian cell-line development. 

The lack of a membrane-barrier in the OCFS provides the opportunity to express and study proteins that are difficult to express in cell-based systems. Many proteins that can't be readily expressed by 

 expression systems due to poor folding, inclusion body accumulation, or due to toxicity can be expressed in an

The advancements in genomic research and increased numbers of sequenced genomes require expression systems that allow fast production of the proteins under investigation. Cell-free expression systems can provide a useful tool for rapid screening and analysis of protein function, which is important for protein-drug discovery and development. DNA molecules can be amplified, transcribed, and translated in microplate wells and the expressed protein can be assayed immediately (15). Recently, HTS in a cell-free wheat germ system led to the discovery of a novel malaria vaccine candidate (16). Finally, the linear scalability allows proteins identified in display-based selections and HTS to be immediately scaled for production of multiple gram quantities, thus avoiding the delays and challenges of conventional mammalian cell line development. 

Currently, cell-based expression technologies exhibit several limitations with respect to protein production at all phases of the drug discovery and development pipeline. Rapid production of proteins with novel chemical modifications, such as ADCs, are particularly challenging.  

-based cell-free protein synthesis systems, however, provide robust, rapid,and scalable protein production. The 

-based OCFS system, in particular, allows rapid and multiplexed production of various difficult to express proteins and opens the unprecedented ability to explore therapeutics beyond the 20 amino acids that define today's proteins (17). The OCFS, combined with rational protein design and the focused use of libraries of nnAAs, allows for rapid exploration and identification of protein therapeutics, moving  from the exploratory stage to clinical scale-up on an unprecedented, rapid  timescale. 

Growing demand for new and better biopharmaceuticals has led to sophisticated advances in protein synthesis that now allow for: 					

Rapid production of target proteins, including those that are difficult to express in cell-based expression systems

Straightforward scalability of protein expression from HTS to commercial levels

Combinatorial screening of many proteins to identify and optimize drug  candidates

Introduction of site-specific chemical modifications, including nnAAs into proteins to improve pharmacological properties

These new approaches to protein expression will revolutionize the development of biopharmaceuticals, and open up the possibility to create drugs that were previously inaccessible, and even unimaginable until now. 

is vice-president of research, both at Sutro Biopharma, 310 Utah Ave, Suite 150, South San Francisco, CA 94080, 

* To whom all correspondence should be addressed

The authors discuss various approaches and related issues, including production of difficult-to express proteins using cell-free expression systems, scalability of protein expression, and site-specific chemical modifications.

Industrializing Design, Development, and Manufacturing of Therapeutic Proteins

The biopharmaceutical industry relies on change to make progress in a commercial environment that simultaneously demands higher productivity, higher quality, and lower costs (1). Recombinant protein titers have improved from tens of milligrams to more than 10 grams per liter during the past 20 years, and at the same time, batch volumes have increased so that industry faces the real prospect of batch yields exceeding 100 kg of protein in the next decade (2). Over the same period, regulatory demands have become more onerous, and the pressure to reduce costs has increased as more biopharmaceuticals come off patent and overseas manufacturers begin to take an interest in Western markets (3, 4). Indeed, the unthinkable is increasingly becoming inevitable—biopharmaceuticals will at some point be regarded as commodities, and manufacturing on the ton scale will be necessary for certain products that are required in large, repetitive doses, such as topical antibody formulations. 

There is no doubt that progress in the industry has been impressive, but the polished bodywork of bioprocessing hides an engine of despair that groans under the strain of current demands. Most of the increases in productivity that have been achieved in previous decades have resulted from improvements in the upstream production phase, with more efficient bioreactors and better media formulations sharing the limelight with cell lines that are intrinsically more productive due to the development of more effective screening technologies to identify the most productive clones (5). Downstream processing is now routinely found to be the bottleneck in biopharmaceutical manufacturing because its capacity has not kept pace with upstream production (1). In some cases, the lack of downstream processing capacity can seriously affect the profitability of a new pharmaceutical product and even result in its failure. Manufacturers facing shrinking margins and less bountiful product pipelines are therefore looking for opportunities to increase the efficiency of downstream processing without inflating the cost of goods sold.  

Perhaps the major challenge facing the biomanufacturing industry today is that downstream processing has no economy of scale (i.e., higher titers translate linearly into higher manufacturing costs) (6). Looking back, we can almost consider the first 15 years of biomanufacturing as a golden era, where manufacturers had the luxury of using inefficient processes because the product itself was far more important (3). Most biopharmaceuticals were required in small doses, and demand was sufficiently low to allow plenty of slack in the system. It was also pointless investing in process efficiency when any tweaks and modifications would arouse the suspicious eye of regulators. It was better to let sleeping dogs lie and be satisfied with the status quo. In this environment, innovation was considered a burden rather than a bonus. 

Inevitably, this relaxed attitude to process efficiency resulted in an immense amount of waste because up to 50% of product batches failed to meet specifications (3). To address this, FDA ordered that processes be designed with quality attributes taken into account (7, 8). The process was no longer a means to an end to achieve the product, but rather the process became part of the product. As the economic screws began to tighten and demand increased, manufacturers turned to the age-old strategy of scaling up their production to achieve cost savings. It was at this time that industry began to flounder.  

Whereas upstream production can be scaled up almost indefinitely by increasing the productivity of cells growing in a bioreactor, downstream processing has limits imposed by physics and chemistry. Downstream processing is driven by the mass of product. If one makes more product per batch, one needs correspondingly larger volumes of buffer, larger storage tanks, preparation areas, larger filters, and most importantly, larger amounts of chromatography media. For the production of antibodies (where Protein A resin is typically used in the primary capture step), the costs of scaling up are in some cases greater than the extra revenue made possible by the increased upstream productivity. Manufacturers find themselves in the paradoxical situation that there is no longer an economy scale in manufacturing, but rather an economic depression reflecting the physical limits that constrain the size of the apparatus used in separations (e.g. chromatography columns and the associated piping, skids, and buffer reservoirs). So far, the extra demand has been absorbed by contract manufacturers offering their spare capacity to fulfill quotas, but this is a short-term measure that cannot cope with the predicted increases in demand as industry faces hundreds of products in clinical development, all requiring at least pilot-scale manufacture according to GMP (9).  

How can this productivity dilemma be addressed? With constant scaling up no longer a viable approach, the sector must return to its roots and innovate to succeed. Three solutions are currently being considered by manufacturers, all inspired in some way by the more encouraging regulatory landscape that rewards rather than punishes innovation. This article examines several issues: the streamlining of existing processes, the revisiting of simple technology solutions currently employed in the bulk-chemical industry, and the use of innovative technologies from the biopharmaceutical research sector. These innovative technologies have the potential to introduce game-changing processing options into an industry still mired in technologies that were the state of the art 20 years ago. On a cautionary note, however, technologies from the research sector can fail, and the rash adoption of new and untested technology platforms can punish an eager company seeking innovative solutions. This is the new dilemma in downstream processing. 

Streamlining and redesigning existing manufacturing processes

Many processes for biopharmaceutical manufacturing were designed at a time where process efficiency was considered unimportant (3). More recently, manufacturers have sought to increase the efficiency of each unit operation, but they are only now starting to consider redesigning the entire process train to see whether cost savings can be made. The trend toward process streamlining owes a lot to FDA's quality-by-design (QbD) principles, which are derived from the design-of-experiments (DOE) concept. The design space of a manufacturing process is littered with efficiency peaks and troughs, but there is not always a simple path leading to the most efficient process. Therefore, process design incorporating efficiency and quality from first principles involves going back to the drawing board and evaluating the critical attributes that contribute to an efficient process. 

Most companies are applying these principles and actively streamlining their processing strategies wherever possible. Antibodies take center stage because they represent more than half of all biopharmaceutical products in development, and their common properties make it possible for companies to share process efficiency data that are applicable across platforms (10, 11). It is for this reason that antibody manufacturing has benefitted from the development of so-called generic platform processes, which are broadly similar for all antibodies but can be tweaked to match the specific properties of individual products (12).  

Antibody manufacturing provides an excellent example of the application of process redesign and streamlining principles to increase productivity, cut costs, and maintain product quality. Most manufacturers use three chromatography steps for antibody purification, starting with a very expensive Protein A capture step that is placed immediately after clarification, followed by anion exchange (AEX) chromatography in flow-through mode to extract negatively charged contaminants, such as host-cell protein (HCP), endotoxins, host DNA, and leached Protein A. Then, either cation exchange (CEX) chromatography or hydrophobic interaction chromatography (HIC) in retention mode is used to remove positively charged residual contaminants and product related impurities, such as aggregates and degradation products (13). Modern platform processes also serve as orthogonal strategies for virus removal. 

Realizing that no further cost savings could be gained by scaling up the above process, Pfizer explored the design space around the standard process and found that certain modifications could reduce costs considerably without affecting the quality of the antibody (14). The company introduced two types of process modifications, one in which the order of the polishing steps was reversed and another in which different separation technologies were used to increase process capacity (i.e., using membrane absorbers for the flow-through chromatography step and replacing the depth filtration step with continuous centrifugation) (15). These changes increased the efficiency of purification to such an extent that, for some antibody products, the cation-exchange step became unnecessary reducing the process from three columns to two columns or even a single column. Not only did this save the direct costs of column resin and buffers, but also reduced the process time by >45% which doubled the productivity in terms of batch processing (14).  

The capacity crunch in downstream processing has been avoided or overcome in other industries by adopting simple and inexpensive technologies (16). In bulk-chemical, conventional pharmaceutical and food and detergent industries, expensive processing solutions, such as chromatography, would never be considered because the costs of implementation would not be sustainable in these high-volume, low-margin processes. This simple approach could be applied to biopharmaceutical manufacturing. 

Several recent developments suggest that simpler technologies could find a niche in biopharmaceutical manufacturing, particularly in the early processing steps where the complex mixture of particulates and solutes have the most potential to foul expensive membranes and resins (16, 17). Tangential flow microfiltration, depth filtration, and (continuous) centrifugation are the current methods of choice for the clarification of the feed stream, and one or more of these processes may be employed in series to remove larger particulates until finally a polishing depth filter or dead-end filter can be used to remove fines and thus reduce feed stream turbidity (18). Efficient and inexpensive clarification becomes more challenging with higher-titer cell culture processes because these are characterized by a greater cell density and often a longer process time, resulting in a higher solids content, more particle diversity (size and physical properties) and—most challenging of all—a greater proportion of fine particles that escape coarse filtration.  A technology that is widely used in the beverage industry and in wastewater processing is the use flocculants to link small particles together and create aggregates that are easier to remove. Flocculation is achieved using polymers that bind simultaneously to the surfaces of several particles through electrostatic interactions, creating larger particles that may sink under gravity, or may be removed more easily by centrifugation or filtration.  

In the bioprocessing industry, flocculation has been used to help remove whole cells from fermentation broth, but more recently it has also been used to remove fine cell debris and proteins. A simple and inexpensive strategy recently applied in antibody manufacturing is the creation of a calcium phosphate precipitate by adding calcium chloride to a final concentration of 30 mM and then potassium phosphate to a final concentration of 20 mM. Precipitation traps cell debris in larger particles, allowing removal by centrifugation for 10 min at 340 X g and yields a clear supernatant with the recovery of ~95% of the antibody (19). Interestingly, this strategy also removes some soluble host cell proteins and nucleic acids. Flocculation is that it does not introduce any additional impurities to the feed stream because the flocculant is removed along with the aggregated particles. 

Precipitation is widely used as a purification approach in the bulk-chemical industry, and given that precipitation can be induced by simple changes in the environment, such as varying the temperature or pH, increasing the salt concentration (i.e., salting out) or adding organic solvents, it should be easy to apply the same principles in bioprocessing (20). Precipitation has therefore been used to remove soluble impurities from the feed stream during antibody manufacturing, and these solids can then be trapped by filtration or pelleted by centrifugation leaving a clear feed stream relatively enriched for the target protein (20). In an innovative adaptation of this approach, the antibody can be precipitated under mild conditions and recovered from a collected pellet thus removing many contaminants in a single step (21). This is possible because the mild precipitation conditions allow the protein to be redissolved without loss of activity. Several groups have developed methods to precipitate antibodies in large-scale processes, and this method could replace Protein A chromatography in the long term (22, 23). Precipitation methods using 

-octanoic acid are used for the removal of contaminants in at least two industrial antibody manufacturing processes (24, 25). 

In the final purification steps, another traditional technology that is being considered for use in biopharmaceutical manufacturing is crystallization. Crystallization involves the separation of a solute from a supersaturated solution by encouraging the growth of crystals. The crystallization process involves the formation of a regularly structured solid phase, which impedes the incorporation of contaminants or solvent molecules, and therefore yields products of exceptional purity (26). It is this purity which makes crystallization particularly suitable for the preparation of pharmaceutical proteins, coupled with the realization that protein crystals enhance protein stability and provide a useful vehicle for drug delivery (27). Protein crystallization has been developed into a commercial technology for drug stabilization and delivery, and several current manufacturing processes involve crystallization including the production of recombinant insulin, aprotinin, and Apo2L (28). 

Although process redesigns and traditional technologies can contribute to the development of downstream processes, they provide only incremental improvements that marginally increase process efficiency. Incremental or evolutionary technologies have been the mainstay of the bioprocessing industry for the past 20 years, and column chromatography provides one of the best examples of this phenomenon in action (29). However, these slow marginal gains are already beginning to decline, and it is becoming difficult to envisage how sustainable processing can continue without a major injection of downstream-processing capacity. 

One way to address this concern is to embrace genuinely novel technological approaches that change the rules of the game. The fringes of the biopharmaceutical industry are populated by companies that survive on innovation, and some of these innovations are disruptive in the sense that their influence on the industry is unpredictable and could contribute to a radical change in bioprocessing. 

Most technological innovations in bioprocessing have been incremental, but there are several recent examples of disruptive innovations that have challenged the established business model and caused real grass-roots change in the industry. Again, many of these changes have affected upstream productivity first (e.g., disposable bioreactors and buffer/media storage bags), but we are also seeing examples in downstream processing (e.g., the introduction of simulated moving bed chromatography, expanded bed chromatography, monoliths, and membrane adsorbers) (1). These innovations have taken hold in niche markets but are now beginning to adopt mainstream positions. Disposable modules for downstream processing occupy a more mature status in the development cycle (30). The use of disposable filter modules is now an industry standard and where filters first left their footprint, membrane adsorbers are set to follow (31). 

 Disposable membranes adsorbers are beginning to replace traditional chromatography in a number of settings, just as disposable membrane filters replaced steel mesh filters. Indeed, membrane filters have evolved into charged filtration devices that use the principles of both sieving and chemical selection to improve filtration performance, thereby creating a precedent for the use of membrane chromatography in downstream processing.  After a period of inertia, the benefits of membrane chromatography are now fairly well established, and manufacturers are willing to consider them as a genuine alternative to fixed columns rather than a step in the dark (32, 33). In contrast to resin-based flow through processes, membrane chromatography involves the use of thin, synthetic microporous or macroporous membranes stacked in layers within a disposable cartridge (34). The footprint of such devices is much smaller than columns with a similar output. A range of membranes is available with functional groups equivalent to the corresponding resins (e.g., membranes containing activated quaternary ammonium groups for anion exchange, or phenyl groups for hydrophobic interaction chromatography [HIC]), and a relatively new variant also allows salt tolerant interaction chromatography (STIC) in high-salt buffers (35, 36). The availability of STIC membranes is an important and innovative advance in biomanufacturing because even the most recent generation of membrane adsorbers fall short of some manufacturing requirements when challenged with the high-conductivity feed streams often produced in high-titer processes. STIC ensures more flexibility in process design and improves the clearance of host cell proteins and viruses in buffers containing high concentrations of salt (see Table I). These new adsorbers therefore allow polishing to be carried out at higher load densities without an interstitial dilution step after product capture, reducing process time and circumventing the need for additional buffer preparation and holding.  

Table I: Broader polishing operation window with salt-tolerant membrane chromatography. 

Table I demonstrates that Sartobind STIC provides higher binding capacities for BSA, DNA, and model viruses compared with a Q anion exchanger under high salt conditions (150 mM NaCl), thereby increasing the design space for polishing operating conditions.  

In a recent example described by the Italian biopharmaceutical company Philogen, membrane adsorbers were substituted for the flow through and bind-and-elute steps for the polishing of a new monoclonal antibody fusion protein in Phase I–II clinical development, achieving 90% recovery and 99.9% purity (37). The performance benefits of membranes provide value for the user, but the complete elimination of cleaning and validation requirements is often cited as the major advantage because this avoids the costs of the chemicals, personnel, and record-keeping, and more importantly avoids the inevitable process down time while cleaning takes place. Spent modules are simply replaced with prevalidated new ones, available in a range of sizes and configurations for maximum flexibility (32). 

The performance advantage of membranes over resins reflects the transport of solutes to their binding sites mainly by convection, while pore diffusion is minimal (see Figure 1a). Because of these hydrodynamic benefits, membrane adsorbers can operate at much greater flow rates than columns, thereby considerably reducing buffer consumption and shortening the overall process time by up to 100-fold.  The use of membrane adsorbers can be viewed as the equivalent of shortening traditional columns to near zero length to allow large-scale processes to run with only a small pressure drop at high flow rates. For example, polishing with an anion exchange membrane can be conducted with a bed height of 4 mm at flow rates of more than 600 cm/h, and provide a high frontal surface area-to-bed height ratio (see Figure 1b). Small-volume disposable membrane chromatography devices can now handle up to 50 L /min/bar/m

. Even at these high flow rates, the membrane pores provide adequate binding capacity for large molecules such as viruses and DNA, so they can play an important role in the overall viral clearance strategy for antibody purification (38, 39).  

Figure 1: (a) Mechanistic comparison of solute transport in bead resins (left) and membrane adsorbers (right), where thicker arrows represent bulk convection, thinner arrows represent film diffusion and curved arrows represent pore diffusion. (b) Comparison of bed height in columns (left) and membrane adsorbers (right). Using membrane adsorbers is functionally equivalent to shortening columns to near-zero length, resulting in a similarly small pressure drop that allows extremely high flow rates, thereby reducing overall process times up to a 100-fold. In this example, both formats have a 1350 cm2 frontal surface; the column has a bed height of 15 cm; and the membrane adsorber has a bed height of 0.4 cm. The height to frontal surface ratio is approximately 100 for the column and nearer to 3500 for the membrane device. (ALL FIGURES ARE COURTESY OF THE AUTHOR) 

 The flexibility of disposable modules is arguably the most important benefit in the context of the whole process, and this reflects the broad industry perspective that manufacturing flexibility is now perhaps at least as important as capacity considering the large numbers of products in clinical development. Process development can be streamlined and expedited because different modules can be tested in various combinations to arrive quickly at the best overall set of process options, and the absence of cleaning and validation requirements can shorten the time required to develop a finalized process by months or years. The ability to replace each module completely also makes it easier to assemble process trains for new products in existing premises without cross-contamination. The flexibility is most noticeable during scale-up because disposable devices are generally modular and available in various sizes, and scaling up simply involves swapping one module for another with a higher capacity. It is thus apparent that membrane devices can be scaled up with none of the attendant disadvantages of column resins, thereby making the goal of polishing 100-kg batches of antibody entirely possible without oversizing.  

Figure 2: Selection guide for convective media, such as membrane adsorbers. HIC is hydrophobic interaction chromatography. STIC is salt tolerant interaction chromatography. 

Innovations that take into account the current state of the industry as well as potential challenges and demands are likely to be the most successful in the long term. At the same time, technologies borrowed from the edge of research always come with risks that must be evaluated by manufacturers looking at major investments into capacity. The perceived bottleneck in downstream processing can be addressed with lower-risk approaches, such as streamlining current production processes, with moderate-risk approaches, such as introducing technologies that have already proven suitable in other industry settings, or with higher-risk approaches involving the incorporation of novel technologies.  

In several cases, these novel technologies have already proven their credentials in several processes and companies following the paths set by the first adopters, the trailblazers of the industry, can be assured that the technologies involved now have established their credibility. 

The future of biomanufacturing is likely to rely more on innovation and flexibility than on traditional strengths, such as large facilities and the financial muscle to invest in them. Disposable manufacturing is likely to play an increasingly important role as companies maneuver in a crowded market to protect their R&D investments while more and more generics become available. The ability to scale up or down quickly, to switch to new campaigns rapidly and to produce multiple products in the same facility will be a key metric of success. Ultimately, the future of bioprocessing will require industry players to embrace the need to change. 

is vice-president of purification technologies at Sartorius Stedim Biotech GmbH, 

 1. U. Gottschalk, "The Renaissance in Protein Purification" supplement to 

 2. D.L. Hacker, M. De Jesus, and F.M. Wurm, 

 4. W. Berthold, Proceedings of BioManufacturing World 2010 (Shanghai, China, 2010). 

 7. FDA, "Pharmaceutical cGMPs for the 21st Century: A Risk-Based Approach" (Rockville, MD, August 2002). 

 8. FDA, "PAT Guidance for Industry: A Framework for Innovative Pharmaceutical Development, Manufacturing, and Quality Assurance" (Rockville, MD, September 2004). 

 12. U. Gottschalk, "New and Unknown Challenges Facing Biomanufacturing," supplement to

 13. S. Vunnum, G. Vedantham, and B. Hubbard  (2009) "Protein A-based Affinity Chromatography," in 

Process Scale Purification of Antibodies, 

U. Gottschalk, Ed. (Wiley, NY, 2009) pp. 79–102. 

The author reviews the state of downstream processing and considers potential solutions, including the streamlining of full processes and borrowed technologies.

The Future of Downstream Processing

RNA-based therapeutics hold significant potential as promising treatment options for human disease. In the past 20 years, advances in the RNA field have identified several novel RNA-based therapies that are currently under clinical investigation, including antisense oligonucleotides, small interfering RNA (siRNA), and microRNA. By targeting RNA and modulating human biology at the molecular level, these new technologies have allowed drug-discovery efforts to focus on a broad range of disease targets once deemed to be "undruggable."  

Leading RNA biotechnology companies have since expanded the target space and generated multiple clinical candidates characterized by improved target specificity, improved drug safety, and demonstrated efficacy in patients. These companies have traditionally focused on targeting specific genes relevant to the disease indication through the control of protein synthesis at the RNA level. More recently, drug discovery researchers are attempting to regulate entire networks of genes through the modulation of a single microRNA. Targeting microRNAs with either oligonucleotide inhibitors, namely anti-miRs, or miR-mimics (double-stranded oligonucleotides that replace microRNA function), provides a novel class of therapeutics and a unique approach to treating disease by modulating entire biological pathways (see Figure 1).  

Figure 1: The RNA therapeutics opportunity. MicroRNAs represent a new set of drug targets capable of regulating an entire network of related genes. (ALL FIGURES COURTESY OF THE AUTHORS)			

Targeting specific genes using antisense oligonucleotides and siRNA

Antisense oligonucleotides and siRNA have great potential to become mainstream therapeutic entities. This is due, in part, to their high specificity and wide therapeutic target space in the genome. The antisense approach targets a specific gene and interrupts the translation phase of the protein production process by preventing the mRNA from reaching the ribosome (1). Antisense drugs are short (15–23mer) chemically modified nucleotide chains that hybridize to a specific complementary area of mRNA. On hybridization, the mRNA is recognized as a RNA-DNA hybrid and degraded through an RNase H cleavage mechanism and not translated by the ribosome into a functional protein (see Figure 2). By inhibiting the production of proteins involved in disease, antisense drugs can create pharmacologic benefit for patients.  

Figure 2: MicroRNAs are key regulators of the genome. Hybridization of microRNAs (red) to their target seed sequence in mRNAs regulates and directs the expression of an entire network of genes. AGO is Argonaute protein, DGCR8 is DiGeorge critical region 8, miR is microRNA, RISC is RNA induced silencing complex.			

RNA interference (RNAi) is a highly conserved sequence-dependent eukaryotic process for regulating gene expression. Small stretches of double-stranded RNA ranging from 19 to 25 base pairs, and known as siRNA, utilize the RNA induced silencing complex (RISC) pathway to target a specific gene and bind to its homologous mRNA. This results in site-specific mRNA cleavage and protein degradation (see Table I) (2). The presence of the RNAi cellular components, combined with silencing, specificity, and efficacy makes it an attractive mechanism for targeting dysregulated gene expression in human disease.  

Table I: Overview of the current RNA-based drug-discovery platform.			

Targeting gene pathways using microRNA therapeutics

More than 750 microRNAs have been identified to date, regulating an estimated one-third of all human genes (3). Using sophisticated bioinformatics analyses and enhanced detection methodologies, scientists demonstrated that a single microRNA may be capable of regulating hundreds of messenger RNAs that function in the same or related pathways. Because microRNAs have functions in multiple biological pathways, a change in expression or function of microRNAs might give rise to diseases, such as cancer, fibrosis, metabolic disorders and inflammatory disorders. The demonstration that several microRNAs are up-regulated in a particular disease phenotype provides the rationale to use anti-miR technology to restore the balance of normal gene regulation inside the cell (see Table I) (4). 

MicroRNAs are small noncoding RNAs that are approximately 20–25 nucleotides in length. They regulate expression of multiple target genes through sequence-specific hybridization to the 3' untranslated region (UTR) of messenger RNAs and block either translation or direct degradation of their target messenger RNAs (5). MicroRNA genes are expressed in the cell nucleus as a precursor called the primary microRNA which, upon further processing by an enzyme called Drosha, lead to pre-microRNA (see Figure 2). Once exported into the cytoplasm, the pre-microRNA is cleaved by the Dicer enzyme into a 20–25 nucleotide-long double-stranded RNA that is then loaded into RISC. This process is followed by the unwinding of the two RNA strands, the degradation of the passenger strand, and the retention of the mature microRNA. Through the RISC, the microRNA guides and targets messenger RNAs through direct base pairing. The 5' region of microRNA, also known as the "seed" region (nucleotides 1 through 8 or 2 through 9), is the most critical sequence for targeting and function (6). The microRNA target sites, located in the 3' UTR of messenger RNAs, are often imperfectly matched to the microRNA sequence.  

MicroRNAs do not require perfect complementarity for target recognition, so a single microRNA is able to regulate multiple messenger RNAs. Although microRNAs exert subtle effects on each individual messenger RNA target, the combined effect is significant and produces measurable phenotypic results. The ability of microRNAs to influence an entire network of genes involved in a common cellular process provides tremendous therapeutic potential and differs from the specificity of today's drugs, which act on specific cellular targets. MicroRNAs play integral roles in several biological processes, including immune modulation, metabolic control, neuronal development, cell cycle, muscle differentiation, and stem-cell differentiation. Most microRNAs are conserved across multiple animal species, indicating the evolutionary importance of these molecules as modulators of critical biological pathways and processes (3).  

The association of microRNA dysfunction with disease has created enormous potential for selective modulation of microRNAs using anti-miR oligonucleotides, which are rationally designed and chemically modified to enhance target affinity, stability, and tissue uptake. Aberrantly expressed or mutated microRNAs that cause significant changes in critical biological pathways represent potential targets whose selective modulation could alter the course of disease. From a mechanistic view, the inhibition of the microRNA target is based on the specific annealing of the anti-miR (see Figure 3). A stable, high-affinity bond between the anti-miR and the microRNA will compete with binding to the 3' UTR target region.  

Figure 3: Single-stranded oligonucleotide anti-miRs pharmacologically modulate dysregulated microRNAs. The anti-miR oligonucleotide (black) binds and hybridizes to the abnormally expressed microRNA (red), blocking its function within the cell. DGCR8 is DiGeorge critical region, and miR is microRNA.			

Studies by Regulus Therapeutics and others have demonstrated that modulating microRNAs through anti-miR oligonucleotides can effectively regulate biological processes and produce therapeutically beneficial results in murine models of cardiac dysfunction; reducing cancer metastases in murine tumor models; and reducing viral load in the chimpanzee model of hepatitis C virus infection (7,8,9). Most recently, advances in oligonucleotide chemistry have improved potency and stability by modification with novel 2',4'-constrained 2'O-ethyl (cEt) nucleotides (10). The ability to achieve increased inhibitory potency with this next generation of bicyclic nucleic acid chemistry could make a significant positive impact on the design of anti-miR inhibitors for a vast array of microRNA disease targets.  

Up until nearly a decade ago, insufficient 

 stability, limited methods of delivery and tissue distribution of oligonucleotides hampered successful clinical development for several promising oligonucleotide therapeutic agents. As high molecular weight, highly charged polyanionic molecules, oligonucleotides faced many hurdles in reaching their target organ or target cell type. First-generation antisense phosphorothiolated oligodeoxynucleotide clinical candidates administered into the bloodstream had a low affinity for their target, poor stability because of nuclease degradation, unfavorable immunostimulatory properties, and rapid excretion by renal clearance, resulting in shortened half-lives (11). To increase their metabolic stability and tissue half-life, antisense and anti-miR oligonucleotides from second-generation nucleoside chemistries were developed that dramatically altered the pharmacokinetic properties of these molecules (10,12).  

The introduction of chemical modifications such as 2' methoxyethyl (MOE) and 2', 4'-constrained 2'O-ethyl (cEt) into the ribose sugar ring significantly improved both the pharmacokinetic and safety profile of antisense oligonucleotides. Once delivered systemically, these second-generation compounds rapidly partition from the plasma and are taken up by cells of multiple tissues without the need of formulation or a delivery vehicle. Benefitting from nearly 20 years of oligonucleotide chemistry advances at companies such as Isis Pharmaceuticals, leading developers of anti-miR therapies have garnered a tremendous advantage in improved delivery strategies. The high water solubility of anti-miR oligonucleotides due to their polyanionic chemical structure has allowed anti-miR formulation in simple aqueous solutions such as buffered saline (13). The only limiting factor is the viscosity of the solution, which is generally concentration-dependent for single-stranded oligonucleotides (13). This simple anti-miR formulation is in contrast to the requirements for double-stranded siRNA drug delivery, which must fully encapsulate the siRNA in a lipid nanoparticle to systemically deliver its contents to a target tissue (14).  

Figure 4: Similar pattern of tissue distribution for chemically modified anti-miRs in mouse and monkey. Anti-miR oligonucleotide quantitation was performed on tissues by either mass spectrometry analysis or capillary gel electrophoresis. IP is Intraperitoneal, SC is subcutaneous.			

Anti-miR route of administration and tissue distribution

Bioavailability and tissue distribution of anti-miRs have been studied extensively in rodents and nonhuman primates. The preferred route of administration for most therapeutic anti-miR compounds is subcutaneous systemic delivery, because it provides efficient dissemination of the drug to different tissues including the liver, kidney and adipose tissue without the need of a drug delivery system. Additionally, the biodistribution of anti-miRs in multiple animal species following subcutaneous administration provides valuable information regarding organs that may be successfully treated, as well as those organs unlikely to be affected. Multiple studies were performed in mice and monkeys with second generation anti-miR 1 and anti-miR 2 compounds given subcutaneously once weekly over several weeks.  A quantitative analysis of tissues demonstrated broad biodistribution of modified anti-miRs among multiple tissue types including the kidney, liver, lymph nodes, adipose tissue, and spleen, as demonstrated by mass spectrometry analysis (see Figure 4). These organs have been previously shown to be target sites for oligonucleotide distribution after parenteral administration (13).  Additionally, the similar pharmacokinetics and correlated tissue distribution of each anti-miR in different preclinical animal models provide important guidance for selection of different disease indications and may assist in better clinical trial designs with anti-miR therapies (see Figure 5). Effective delivery of anti-miR oligonucleotides has also been demonstrated in different species through multiple routes of administration including: intravenous, intraperitoneal, intratracheal, intranasal, and intracerebral. A more detailed analysis of anti-miR tissue distribution using quantitative whole body autoradiography to provide additional quantitative information is in progress.  

Figure 5: The distributions of anti-miRs in mice and monkey are highly correlated. Quantitative analysis of drug concentration by mass spectrometry revealed a good correlation of drug tissue distribution across multiple species.			

mRNA expression profiling methods coupled with statistical techniques that can measure small changes in the expression of many genes have become powerful tools to further our understanding of the biological role and function of microRNAs. Relying on the scientific findings that some microRNAs are capable of regulating hundreds of messenger RNAs, studies were performed in mice to determine anti-miR delivery to different cell types. Mice were treated with a specific anti-miR (intraperitoneal injection) and multiple cell types were harvested to for mRNA expression studies using Sylamer enrichment analysis (15). Anti-miR oligonucleotides are distributed to peritoneal macrophages as evidenced by flow cytometry analysis and target gene up-regulation (see Figure 6). An analysis identifying an overrepresented set of genes associated with a specific anti-miR biological effect was conducted and a data plot from the isolated macrophages was generated that demonstrated the most up-regulated sets of genes after anti-miR treatment. 

 values generated for this dataset suggest statistically significant preferential up-regulation of genes matched to their target sequence after anti-miR treatment.  

Figure 6: Functional drug delivery of anti-miRs in mouse peritoneal macrophages. Flow-cytometry studies (a) and gene-regulation studies (b) demonstrate the internalization of anti-miR and target engagement in macrophages (Sylamer analysis). Potential targets containing heptamer 1-7 (GCATTAA) or heptamer 2-8 (AGCATTA) are enriched. X-axis is ranked genes by fold change. Y axis is -log (P-value enrichment). PBS is Phosphate-buffered saline.			

Targeting pathways of human disease with microRNA-based drugs represents a novel and potentially powerful therapeutic approach. Recent data demonstrate not only that dysregulated microRNAs are associated with and can cause human disease, but that selective modulation through anti-miR intervention can provide therapeutic benefits. Anti-miR oligonucleotides can be easily administered through local or parenteral injection routes with sufficient uptake of the agent to achieve sustained target inhibition in tissues and organs without the need of formulation. Improvements in anti-miR chemical design and pharmacokinetic properties will allow further exploration of microRNA biology and broaden the utility of microRNA therapeutics. 

, is the associate director of chemistry, and 

, is the senior director of chemistry, all at Regulus Therapeutics, 3545 John Hopkins Ct., San Diego, CA 92121, tel. 858.202.6321, 

*To whom all correspondence should be addressed. 

 1.  S.T. Crooke et al., "Mechanisms of Antisense Drug Action, an Introduction," in 

Antisense Drug Technology: Principles, Strategies, and Applications,

 S.T. Crooke, Ed. (CRC Press, Boca Raton, 2007), pp. 3–47. 

 11. A.A. Levin, R. Zu, and R.S. Geary, "Basic Principles of the Pharmacokinetics of Antisense Oligonucleotide Drugs," in 

 S.T. Crooke, Ed. (CRC Press, Boca Raton, 2007), pp. 183–215. 

 13. R.S. Geary, R. Zu, and A.A. Levin, "Pharmacokinetic/Pharmacodynamic Properties of Phosphorothioate 2'-O-(2-Methoxyethyl)-Modified Antisense Oligonucleotides in Animals and Man," in 

 S.T. Crooke, Ed. (CRC Press, Boca Raton, 2007), pp. 306–326. 

The authors provide further insight into microRNA biology, and the simplicity of anti-miR oligonucleotide drug delivery.

Perspectives in MicroRNA Therapeutics

This article is part of a special issue on 

Biopharmaceuticals continue to be the hot growth sector, and most revenue projections suggest that biopharmaceuticals will occupy an increasing number of spots in the top 10 highest revenue generators in coming years. Nevertheless, the results from the third annual 

Bioprocessing survey indicate a slowdown in new entrants to the market. In 2009, more than 50% of respondents said they were entering the biopharmaceutical manufacturing arena for the first time. In 2010, only 23% did. But new entrants and veterans alike all face significant challenges and opportunities, and 249 respondents, representing firms large and small told us about them throughout March 2011. 

Just as in years past, slightly under half of respondents (48%) produce biopharmaceuticals only, with the remainder working for companies that produce both traditional small-molecule pharmaceuticals as well as biopharmaceuticals. As in the past, there is a dramatic division in company revenues—the majority of respondents producing biopharmaceuticals only work at companies with revenues under $10 billion; the majority of respondents producing both biopharmaceuticals and traditional small-molecule drugs work for companies with revenues above $10 billion. Half of all respondents added biopharmaceutical manufacturing capacity this year, but they come from companies of all revenue classes, with 86% of respondents from companies with revenues up to $1 billion and 88% of respondents from companies with revenues more than $50 billion saying their companies increased capacity. Fifty-seven percent of respondents increased capacity because they added new products to their line (about 72% of those were products developed internally), and 52% increased production of an existing product. Approximately 18% of respondents acquired a company with a biomanufacturing program, and an equal number in-licensed a biopharmaceutical product that necessitated a capacity increase. Among those who decreased their capacity, 31% decreased production of an existing product, and an equal number outsourced production of a product that had previously been manufactured in house.  

Figure 1: What class(es) of biopharmaceutical does your company manufacture? (multiple responses allowed) 

Therapeutic proteins other than monoclonal antibodies (mAbs) continue to constitute the largest product class, produced by 51% of respondents, followed by monoclonal antibodies (45%), and vaccines (29%). These percentages reflect some interesting changes from last year's product mix. The percentage of respondents involved in monoclonal antibody production is up slightly, about five percent over last year, while vaccine manufacturing is down about seven percent. The percentage of respondents involved in nucleic acid products remained virtually unchanged—14% this year, versus  12% last year, while there was a slight uptick in the number of respondents producing cells for cell and tissue therapies, 14% this year compared with 10% last year. Forty-five percent of respondents say their companies plan to produce follow-on biologics, up slightly from last year's 41%. The remainder either will not (21%) or don't know (34%). Eleven percent of respondents say their companies produce only follow-on biologics right now, while 23% say they  produce both follow-on and branded products.  

Among those who manufacture proteins, producing adequate yields and purifying them remain the two most vexing problems, with an equal number—41%—of respondents identifying these as challenges. However, if we look closer, we find that 47% of those producing non-mAb proteins saying that purifying product is a challenge (versus 40% who produce mAbs). About equal numbers (45% of mAb producers; 43% of non-mAb producers) say producing high enough yields is a challenge. In general, the challenges in protein production are similar regardless of the type of protein produced, with about 44% of respondents of both classes of proteins saying they have problems with protein aggregates, and a slightly lower percentage expressing concern over protein stability.  

Figure 2: What kind of equipment do yo uhave in  your biopharmaceutical manufacturing facility right now? 

Purification and stability are also issues for manufacturers of nucleic acids, with 47% of respondents identifying each of these as key challenges. At 33%, scale up is also a challenge for nucleic acid manufacturers. Process development is the major challenge for 43% of producers of cells for cell and tissue therapies, followed by cell stability (36%) and process scale up (34%). 

The debate continues over the use of disposables in biopharmaceutical manufacturing, with firms considering the convenience, cost, and the environmental impact of single-use, largely plastic, components. Anecdotally, it seems the industry is increasingly embracing disposable components. Yet our survey indicates an insignificant increase in the number of respondents using all-disposable systems—9% this year versus 8% last. Hybrid systems, which combine stainless and disposable components, remains the most popular choice, but the 66% of respondents using hybrid systems represents a notable decline from last year's 72%, which was down from the previous year's 74%. The use of stainless components continues to rise—26% of respondents this year report using all stainless equipment, an increase over last year's 20%. Interestingly, 11% of respondents who now use hybrid systems say they plan to convert to fully disposable equipment in the coming year, while no respondents anticipate moving to an all-stainless system. No companies with revenues above $5 billion use completely disposable systems; 88% of respondents from companies with revenues in excess of $50 billion use hybrid systems.  

Table I: What is (are) the greatest advantages to disposable equipment? (Multiple answers allowed) 

As in years past, we detected some marked differences in perceived advantages and challenges with disposable equipment among users of that equipment versus non-users. The first discrepancy arises over the question of cost savings. Thirty-five percent of all-stainless users (and 31% of hybrid users) believe there's a cost advantage to all-disposable systems. In contrast, 54% of all-disposable users say cost is an advantage to all-disposable systems, suggesting users of the other two systems underestimate the savings to be gained by an all-disposable system. Those who use all-disposable systems also perceive disposables to be more environmentally friendly (23%) than do all-stainless users (15%) and hybrid users (10%). All-stainless users also underestimate process consistency that can be achieved with all-disposable equipment, as only 18% of stainless users cite this as an advantage, contrasted with 31%, who actually use all-disposable systems. Additional responses suggest it may be easier to incorporate automation and may have greater regulatory acceptance than all stainless users might think. 

These same discrepancies arise when respondents were asked about the challenges inherent in single-use technologies. Users of all-stainless equipment overestimated the cost, risk of contamination, and process inconsistency, compared with respondents who use all-disposable equipment. On the other hand, all-stainless users underestimated the difficulty of incorporating process analytical technologies, implementing QbD principles, and the risk posed by extractables and leachables. In addition, users of hybrid technologies reported having greater challenges gaining regulatory acceptance (12.4%) than did users of all-disposable equipment, none of whom (0%) reported this as a challenge.  

Table II: What is (are) the greatest challenges to disposable equipment? (Multiple answers allowed) 

The percentage of respondents who say their companies outsource the manufacture of biologics rose again slightly this year over last year (36% this year versus 33% last), which represented a small rise over the previous year (30%). The reasons for outsourcing seemed to have changed slightly from last year's reasons. This year, 51% of respondents say their companies outsource to compensate for no or limited manufacturing capacity, whereas only 45% gave this reason last year. The percentage citing cost effectiveness climbed this year, to 49%, up from 41% last year. Companies with revenues under $10 billion are most likely to outsource, and their reasons for doing so differ, as well. A greater percentage of respondents from companies with revenues under $5 billion outsource because they have no or limited manufacturing capacity (for example, 71% of respondents from companies with revenues between $1 billion and $5 billion give this as the reason). Companies with revenues greater than $5 billion outsource to realize cost savings. Smaller companies—those with revenues under $500 million also say they lack the expertise internally (up to 75% of respondents from companies with revenues under $250 million, as opposed to 33% from companies with revenues between $5 billion and $10 billion). 

Overall, 61% of respondents say they outsource the manufacture of clinical-trial materials, and 49% said they outsource the manufacture of both drug substance (APIs) and finished drug product. Thirty-three percent outsource the manufacture of drug substance only, and an equal percentage (31%) said they outsource process development and full commercial-scale manufacture. The functions least likely to be outsourced were drug product only, and formulation development (29% each). There were no clear-cut patterns when it came to the size of companies and the functions they outsourced. In general, companies with revenues under $500 million were more likely to outsource process and formulation development, than were companies with larger revenues. Whereas companies with revenues above $500 million were more likely to outsource the manufacture of both drug substance and drug product than were smaller companies.  

Table III: Challenges according to protein class. 

There was no appreciable change over last year in the number of respondents reporting that their companies incorporate quality-by-design (QbD) principles into their operations—65% this year versus 67% last. Nor is there a significant difference over last year in the percentages of respondents who say QbD helps to improve their process understanding (69%), manufacturing efficiency (50%), and improved product quality (49%), while reducing variability (47%). Among those who do not incorporate QbD into their processes, 42% say they lack sufficient guidance from regulatory agencies, 25% say they don't understand the initiative, and 23% don't see any advantages to be gained. Finally, 21% say QbD is too costly. Companies with revenues above $1 billion are more likely to incorporate QbD than are smaller companies—with 92% of respondents in companies with revenues between $10 billion and $50 billion, and 100% of respondents in companies with revenues above $50 billion saying they incorporate QbD. In contrast, only 50% of respondents in companies with revenues under $500 million say they do. Respondents in smaller companies are less likely to understand the initiative and more likely to find it too costly than are their counterparts in companies with larger revenues. 

While 65% of respondents say they incorporate QbD into their manufacturing operations, only 44% say they incorporate process analytical technology (PAT), although here there is a dramatic difference between larger and smaller companies. Fewer than 60% of respondents from companies with revenues below $5 billion say they incorporate PAT, whereas more than 70% of respondents from companies with revenues higher than $5 billion say they do.  High performance liquid chromatography (HPLC) is far and away the most frequently employed analytical technology, with 69% of respondents saying they use it. That's followed by 31%  each who say they use disposable sensors and Fourier-transform infrared spectroscopy (FT-IR).  

As the economy comes out of the recession, the hope among equipment manufacturers is that customers will increase their spending in 2011, compared with 2010. Will they? Interestingly, only 26% say they plan on increasing their equipment purchases this year over last (down from 31% who made that prediction last year—a very accurate prediction, since exactly that percentage report that they did in fact increase their spending levels in 2010 over 2009 levels). Another 36% say they expect spending levels in 2011 to remain the same as 2010, and 13% (down from last year's 15%) say they expect to decrease their spending levels. 

Finally, we asked respondents to tell us what innovations would improve their process efficiency and/or product quality. This year, respondents hoped for better automation, better control of post-translational modifications and analytics that can discern small variations in protein composition (in real time) and in cell metabolism. Others hoped for more streamlined purification processes and an alternative to Protein A. 

Results from our annual survey. This article contains online bonus material and is part of a special issue on Bioprocessing and Sterile Manufacturing.

Pharmaceutical Technology-05-01-2011

Pharmaceutical Technology North America, Bioprocessing Supplement 2011 (PDF)

Methods for the Automated Manufacturing of an Autologous Dendritic-Cell Immunotherapy

Industrializing Design, Development, and Manufacturing of Therapeutic Proteins

The Future of Downstream Processing

Perspectives in MicroRNA Therapeutics

Bioprocessing Trends: Annual Survey

Regulation of Aseptic Processing in the 21st Century

Technologies for Downstream Processing in Biologics

Peptide PEGylation: The Next Generation

Strategies in Parenteral Drug Manufacturing