Development of a Novel Rolling-Tube Cell Culture Platform and Demonstration of System Feasibility - A new rolling-tube system balances scale-up accuracy and thoroughput. - BioPharm International


Development of a Novel Rolling-Tube Cell Culture Platform and Demonstration of System Feasibility
A new rolling-tube system balances scale-up accuracy and thoroughput.

BioPharm International
Volume 23, Issue 8


The rolling-tube platform combines two low cost, off-the-shelf products to create a scaled-down roller bottle platform. The platform comprises 50-mL tubes with vented caps, allowing for superior gas exchange (available from TPP and Sartorius) placed on a rolling device with speed control. These tubes (trade name Cultiflask 50) have been used to grow cells by mounting them vertically on an orbital shaker platform.2,3 Mixing using a rolling motion provides good aeration but low shear, which is important for fragile, agitation-sensitive cells, and certain media formulations. The tubes are angled to prevent the media from wetting the filter on the cap, and the rolling device is placed inside a humidified, temperature- and CO2-controlled incubator. The device can operate continuously inside the incubator. The rolling-tube platform can operate at a throughput of 16 tubes per platform, and because the device is stackable, it requires a small footprint. The 50-mL growth vessel also can serve as the harvest vessel, facilitating media exchanges (e.g., during cell splitting) or cell harvest through standard centrifugation. The current set up allows for fill volumes of up to ~14 mL. This volume may be increased in a future improvement of tube design by centralizing the vents on the tube's cap. The rolling device may be improved as well. The current model is available as a "rock and roll" model. We had to request that the rocking motion be eliminated, requiring customization by the manufacturer to a "roll-only" model to prevent filter wetting during culture.


Proprietary commercial suspension-adapted mammalian production cells and media were used throughout the study. Incubator settings (temperature, % CO2, and humidity) were set to correspond to commercial manufacturing settings used in perfusion bioreactors. Intervals between cell passaging and target seeding density were optimized to correspond to the cell-specific perfusion rate of the production-scale bioreactors. Data are presented for a four-day batch culture. Up to 16 vented-cap 50-mL tubes can be placed on the 9-roller platform that was operated at 30 rpm. No significant difference in culture performance was observed up to the maximum speed of 55 rpm. T-75 flasks and 6-well plates, both nontissue culture treated to prevent adhesion of the suspension-adapted cells, were placed on a standard orbital shaker operated at 35 rpm.

Both the rolling platform and shaker were placed in the same incubator. A blood gas analyzer (BGA) was used to determine pH, pCO2, and pO2. Cell count, viability, nutrients, metabolites, and osmolality were determined using a NOVA BioFLEX instrument. Daily samples of 1.3 mL were taken for BGA and NOVA analysis. Potency (also used for specific productivity calculation) was determined using a product-specific assay. To set up the experiments, cells passaged and scaled up using T-75 flasks were used to inoculate the three platforms at 0.35 x 106 viable cells/mL, as follows: two T-75 flasks (duplicates) with 20 mL working volume, two 6-well plates with 2.9 mL working volume/well, and five 50-mL rolling tubes (two duplicates and three for individual sampling) with 14-mL working volume. This setting was meant to test robustness of the rolling tube platform across different working volumes. One set decreased in working volume along the culture days because of daily BGA and NOVA sampling, whereas in the other set of tubes, one tube was sampled each day, so the working volume remained constant.

Figure 1. On each of the four days of the study, samples were taken from: A) the same 50-mL rolling tubes (volume reduced along study), B) a single 50 mL rolling tube sacrificed for sampling (constant volume along study), C) T-flasks, and D) 6-well plates. Error bars represent the average of two samples (duplicates). The same cell pool was used to seed the cultures on day 0, therefore the values are the same across all platforms. A and B values for day 1 are the same because the tube sampling regimens were shared at the first sampling point.
To determine the effect on culture performance in the rolling tubes of volume reduction caused by daily sampling, (considering the greater percentage change in volume when sampling from small working volume platforms) three rolling tubes were included to be sampled only once, each on a different day, throughout the four-day batch culture. In this set, a 1.3-mL sample was taken each day from a different tube on day 2, 3, or 4. Potency samples (1.5 mL) were taken on days 3 and 4 for each culture to test for productivity.

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