Optimization of bioprocess conditions improves production of a CHO cell-derived, bioengineered heparin.

Heparin is the most widely used anticoagulant drug in the world today. Heparin is currently produced from animal tissues, primarily porcine intestines. A recent contamination crisis motivated development of a non-animal-derived source of this critical drug. We hypothesized that Chinese hamster ovary (CHO) cells could be metabolically engineered to produce a bioengineered heparin, equivalent to current pharmaceutical heparin. We previously engineered CHO-S cells to overexpress two exogenous enzymes from the heparin/heparan sulfate biosynthetic pathway, increasing the anticoagulant activity ∼100-fold and the heparin/heparan sulfate yield ∼10-fold. Here, we explored the effects of bioprocess parameters on the yield and anticoagulant activity of the bioengineered GAGs. Fed-batch shaker-flask studies using a proprietary, chemically-defined feed, resulted in ∼two-fold increase in integrated viable cell density and a 70% increase in specific productivity, resulting in nearly three-fold increase in product titer. Transferring the process to a stirred-tank bioreactor increased the productivity further, yielding a final product concentration of ∼90 µg/mL. Unfortunately, the product composition still differs from pharmaceutical heparin, suggesting that additional metabolic engineering will be required. However, these studies clearly demonstrate bioprocess optimization, in parallel with metabolic engineering refinements, will play a substantial role in developing a bioengineered heparin to replace the current animal-derived drug.

Screening and optimization of chemically defined media and feeds with integrated and statistical approaches.

The majority of therapeutic proteins are expressed in mammalian cells, predominantly in Chinese Hamster Ovary cells. While cell culture media and feed supplements are crucial to protein productivity, medium optimization can be labor intensive and time-consuming. In this chapter, we describe some basic concepts in medium development and introduce a rational and rapid workflow to screen and optimize media and feeds. The major goal of medium screening is to select a base formulation as the foundation for further optimization, but ironically, the most conventional screening method may actually rule out ideal chemically defined medium candidates. Appropriate cell adaptation is the key to identifying an optimal base medium, particularly when cells were originally cultured in serum-free medium containing recombinant proteins and/or undefined hydrolysates. The efficient workflow described herein integrates the optimization of both medium and feed simultaneously using a Design-of-Experiment (DOE) approach. The feasibility of the workflow is then demonstrated with a case study, in which chemically defined medium and feed were optimized in a single fed-batch study using a high-throughput microbioreactor system (SimCell™), which resulted in improving protein titers three- to sixfold.

Identifying the CHO secretome using mucin-type O-linked glycosylation and click-chemistry.

Chinese hamster ovary cells (CHO) are the most common cell line used in the production of therapeutic proteins. Understanding the complex pattern of secreted host cell proteins (HCP) that are released by CHO cells will facilitate the development of new recombinant protein production processes. In this study, we have adapted the N-azido-galactosamine (GalNAz) metabolic labeling method to enable the mass spectrometry identification and quantification of secreted proteins in cell culture media. CHO DG44 and CHO-S cells were cultured in media containing GalNAz, which was metabolically incorporated into mucin-type O-linked glycans of secreted proteins. These proteins were effectively enriched using click-chemistry from the cell culture media, allowing for the analysis of secreted proteins across multiple days of cell growth. When compared to the standard method for secretome analysis, the GalNAz method not only increased the total number of proteins identified but dramatically improved the quality of data by decreasing the number of background proteins (cytosolic or nuclear) to essentially zero.

An HPLC-MALDI MS method for N-glycan analyses using smaller size samples: application to monitor glycan modulation by medium conditions.

Existing HPLC methods can provide detailed structure and isomeric information, but are often slow and require large initial sample sizes. In this study, a previously established two-dimensional HPLC technique was adapted to a two-step identification method for smaller sample sizes. After cleavage from proteins, purification, and fluorescent labeling, glycans were analyzed on a 2-mm reverse phase HPLC column on a conventional HPLC and spotted onto a MALDI-TOF MS plate using an automated plate spotter to determine molecular weights. A direct correlation was found for 25 neutral oligosaccharides between the 2-mm Shim-Pack VP-ODS HPLC column (Shimadzu) and the 6-mm CLC-ODS column (Shimadzu) of the standard two- and three-dimensional methods. The increased throughput adaptations allowed a 100-fold reduction in required amounts of starting protein. The entire process can be carried out in 2-3 days for a large number of samples as compared to 1-2 weeks per sample for previous two-dimensional HPLC methods. The modified method was verified by identifying N-glycan structures, including specifying two different galactosylated positional isomers, of an IgG antibody from human sera samples. Analysis of tissue plasminogen activator (t-PA) from CHO cell cultures under varying culture conditions illustrated how the method can identify changes in oligosaccharide structure in the presence of different media environments. Raising glutamine concentrations or adding ammonia directly to the culture led to decreased galactosylation, while substituting GlutaMAX-I, a dipeptide of L-alanine and L-glutamine, resulted in structures with more galactosylation. This modified system will enable glycoprofiling of smaller glycoprotein samples in a shorter time period and allow a more rapid evaluation of the effects of culture conditions on expressed protein glycosylation.