Name: primary clarification of cho harvested cell culture fluid using an acoustic separator
Uploaded: 2020-05-14T13:30:00.000+00:00
Duration: 7 min 6 s
Description: Watch this Scientific Journal Video about Primary Clarification of CHO Harvested Cell Culture Fluid using an Acoustic Separator at JoVE.com

The authors would like to acknowledge Nilou Sarah Arden and Zhong Zhao for their constructive review of this manuscript. The authors would also like to thank Lindsey Brown for essential input during this project. Partial internal funding and support for this work was provided by the CDER Critical Path Program (CA #1-13) and the CDER Manufacturing Science, Innovation Center of Excellence Program (Berilla-CoE-19-49). This project was supported in part by the Internship/Research Participation Program at the Office of Biotechnology Products, U.S. Food and Drug Administration, administered by the Oak Ridge Institute for Science and Education through an interagency agreement between the U.S. Department of Energy and FDA.
This publication reflects the views of the author and should not be construed to represent FDA&#8217;s views or policies.

1. Preparation of AWS
NOTE: This protocol was developed using one acoustophoretic chamber. However, the bench-scale AWS has five turbidity probes that can operate 4 acoustophoretic chambers in series if needed.
<ol>
	<li>Connect the turbidity cables into their respective ports labeled as F, 1, 2, 3, 4 (e.g., turbidity probe 1 in Figure 1c and port 1 in Figure 2a) and the chamber power BNC cables to the back of the AWS system labeled as 1, 2, 3, 4 (Figure 2b). 
	NOTE: Do not connect the other end of the chamber power BNC cable (Figure 3c) to the back of the acoustophoretic chamber (Figure 3d) until step 2.6 when the chamber is filled with fluid. If the chamber power is turned on with no fluid in the chamber, it will damage the piezo transducer in the chamber and no longer function.</li>
	<li>Insert the turbidity probes into the turbidity meter and thermometer housing and tighten the screws (Figure 1c and Figure 3).</li>
	<li>Connect the feed tubing to the input of the feed turbidity port (Figure 4a) via the feed pump (Figure 1b to 1c).</li>
	<li>Connect the y-tubing from the output of the feed turbidity port (Figure 4b) to the inlet ports of the acoustophoretic chamber (Figure 4c).</li>
	<li>Connect the stage1 tubing from the waste port of the acoustophoretic chamber (Figure 4d) via the stage1 pump to a cell collection vessel (Figure 1d via 1e to 1f).</li>
	<li>Connect the tubing from the permeate port of the acoustophoretic chamber (Figure 4e) to the input of probe1 turbidity port (Figure 4f).</li>
	<li>Connect the harvest tubing from out of the probe1 turbidity port (Figure 4g) to a product collection vessel (Figure 1c to 1g).</li>
</ol>
2. Prime the system with HCCF
NOTE: This protocol was developed using CHO-K1 cells cultured in chemically-defined medium (ActiCHO P with 6 mM L-glutamine) producing the model IgG1 monoclonal antibody VRC0113 and may need to be adjusted for other cell lines and products. The HCCF used in this protocol was obtained at the end of 7&#8211;8 days of CHO-K1 cultures from a shaking flask or bioreactor process.
<ol>
	<li>Turn on the AWS by turning on the power switches on the back and front of the AWS.</li>
	<li>Turn on the computer and double-click the desk icon for the associated software (see Table of Materials). From the &#8220;Readings&#8221; panel, press the &#8220;Start Test&#8221; button to initiate data recording (Figure 5). Once data recording is initiated, all data collected will be recorded in an exportable spreadsheet until the test is stopped.</li>
	<li>Connect the feed tubing end into the HCCF vessel being stirred.</li>
	<li>Start the feed pump by entering the pump rate on the &#8220;Controls&#8221; screen within the &#8220;Feed Pump Image&#8221; and press &#8220;Enter&#8221; on the keyboard. Ensure that the &#8220;Pump Direction Arrow Icon&#8221; (clockwise or counterclockwise) is correctly selected in the gray box to the right of the Feed Pump Image to pump HCCF from the vessel into the acoustophoretic chamber. Click on the &#8220;Triangle Icon&#8221; next to the words &#8220;Turn ON&#8221; within the gray box under the Feed Pump Image to &#8220;Start&#8221; the pump (Figure 6a). 
	NOTE: The maximum pump rate is 10 L/h (167 mL/min), but it is recommended that the nominal flow rate, 60 mL/min, be used for this step to quickly fill the tubing and chamber without risking overfilling the chamber prior to beginning the separation.</li>
	<li>Monitor the feed turbidity measurements by observing the &#8220;Percent Reduction Panel&#8221; (Figure 5c) during the filling of the acoustophoretic chamber. The turbidity values will remain consistent during the loading of the chamber if the HCCF is being mixed sufficiently in the HCCF vessel.</li>
	<li>Once the liquid is above the piezo transducer at the back of the acoustophoretic chamber, press &#8220;Turn OFF&#8221; within the gray box under the Feed Pump Image to stop the pump. Connect the BNC power cable (Figure 3c) to the acoustophoretic chamber (Figure 3d). 
	NOTE: The holdup volume of each acoustophoretic chamber is around 190 mL.</li>
</ol>
3. Operation of AWS
<ol>
	<li>Once the acoustophoretic chamber is filled and the BNC power cable is connected to the back of the acoustophoretic chamber, change the rate of feed pump to the desired operating rate (Figure 6a and Table 1). Several feed pump rates should be tested to identify the ideal operating parameters for a given process.</li>
	<li>Turn on the stage1 piezo power by sliding the bar in the power module to 10 W (Figure 6d) and pressing the &#8220;Turn ON&#8221; icon on the right side of the Stage 1 box (Figure 6c). As suggested by the manufacturer, use 10 W, the recommended power setting for CHO cells. This will generate a fixed frequency of 2 MHz for operation. After several seconds, the cells will start to visibly clump at the wave nodes in the acoustophoretic chamber (Figure 7a).</li>
	<li>Once the cells begin to settle to the bottom of the acoustophoretic chamber (Figure 7b), start the stage1 pump with an appropriate rate based on the cell density and feed pump rate (Figure 6b). Based on the manufacturer&#8217;s optimization and steady-state operation spreadsheet, the stage1 pump rate can be calculated based on the packed cell mass and feed flow rate using the following equation 
	<img alt="Equation 1" src="/files/ftp_upload/61161/61161equ01.jpg" />
	<ol>
		<li>To calculate packed cell mass, tare a scale with an empty 15 mL tube (or other tube compatible with centrifugation). Fill the tube with feed material and record the total weight of the tube with feed. Centrifuge the tube for 10 min at 3,700 x g. Decant the supernatant into a separate container. Measure the weight of the tube with the cell pellet. The packed cell mass percentage of the feed material = (decanted tube weight/filled tube weight) x 100%.</li>
	</ol>
	</li>
	<li>Monitor the turbidity profile of stage1 turbidity as the overflow from the acoustophoretic chamber enters the turbidity probe1 (Figure 8).</li>
	<li>As the cell separation continues, cell removal efficiency <img alt="Equation 2" src="/files/ftp_upload/61161/61161equ02.jpg" /> will increase.</li>
	<li>In addition, monitor the temperature difference between feed and stage1 as it will increase as the cell separation continues (Figure 9). The temperature may rise when higher cell density feed is used but can generally be reduced by altering the feed flow rate. 
	NOTE: When using multiple acoustophoretic chambers, one can sequentially connect the tubing from the previous acoustophoretic chamber via turbidity probes to the input of the current acoustophoretic chamber. In addition, one can connect stage tubing from the bottom of the acoustophoretic chambers via stage pumps to cell collection vessels. A total of four acoustophoretic chambers can be connected in series for optimal cell removal efficiency if necessary.</li>
</ol>
4. Ending AWS
<ol>
	<li>When the run is over, stop the feed and stage1 pump by pressing Turn OFF within the gray box under the feed pump and stage1 pump images, respectively, to stop the pumps.</li>
	<li>Turn off the power to the chamber by pressing Turn OFF on the right side of the Stage 1 box and disconnect the BNC power cable.</li>
	<li>Take the product harvest material collected for further clarification and purification procedures, such as depth filtration and chromatography methods. Discard the cell harvest material.</li>
	<li>Drain the remaining fluid in the acoustophoretic chamber by placing the waste tubing into an empty vessel, disconnecting the tubing from the acoustophoretic chamber inlet and permeate ports and releasing the waste tubing from the pump head.</li>
	<li>Reconnect the tubing, place the waste tubing back into the pump head, and flow through DI water from the end of feed tubing using a feed pump rate of 60 mL/min and stage1 pump rate of 60 mL/min. Continue for 15&#8211;20 min. Discard the flow through.</li>
	<li>Pump 70% isopropyl alcohol (IPA) through the tubing and the chamber using the feed pump for 15&#8211;20 min. Discard the flow through.</li>
	<li>Repeat cleaning procedure with DI water to clear the tubes and the chamber by using the feed pump for 15&#8211;20 min. Discard the flow through.</li>
	<li>Disassemble the tubing from the turbidity probes and acoustophoretic chambers and clean the area with 70% IPA.</li>
	<li>Disassemble the turbidity probes and clean inside the turbidity probes with 70% IPA. Let all parts air dry and then reassemble for the next use. 
	NOTE: The acoustophoretic chambers are manufactured for single use but can be reused if handled and cleaned properly as described in this protocol.</li>
</ol>

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The authors have no financial interests in the products described in this manuscript and have nothing else to disclose.

Described is a step-by-step protocol for the implementation of a bench-scale AWS in primary clarification of a model monoclonal antibody from HCCF of a CHO cell line. As shown in the representative results, the use of the AWS in primary clarification resulted in effective cell clarification and product recovery. Furthermore, the low level of maintenance and operational requirements and scale-up capability allow broader application potential in primary clarification.
Importantly, the representative results suggest that the feed pump rate is critical for the separation of the cells. In addition, due to limitations in detecting high cell density in the turbidity measurement, working cell density is another factor to consider when using an AWS system. Because the turbidity probe will be saturated when running high cell density feed material, it may be best to calculate cell separation efficiency by measuring the cell densities of the feed material and clarified cell culture fluid offline. With accurate offline measurement of clarification, one process strategy that can be considered in solving these problems is to use multiple chambers in series. Although this protocol is primarily focused on using a single chamber, this system can operate three additional chambers for sequential clarification of the cells that can minimally impact the condition of the cells and result in high product recovery. In addition, other parameters, such as power for AWS and cell removal flow rates, can be further optimized for specific cell types or mode of operation. Overall, an optimization of the operation parameters and strategy using these considerations are recommended prior to implementation of AWS.
Among many application potentials, the use of AWS in continuous bioprocessing is promising. Because AWS can replace centrifugation and drastically reduce the filter surface area14, the use of AWS would enable a constant flow of cell-free material for subsequent filtration and chromatography processes that are compatible with continuous biomanufacturing. Due to this compatibility and the availability of perfusion technology for high cell density (e.g., &#62;50 x 106 cells/mL) and longer culture duration (e.g., &#62;14 days), AWS has the potential to develop a novel continuous cell bleeding strategy in addition to the primary clarification.
In some cases, up to 30% of the protein therapeutic produced is removed during steady-state operation in the cell bleed material15,16. In addition, for the removed monoclonal antibodies to be pooled with the standard harvested material certain quality attributes must be met. When these specifications are not met, the result may be a rejection of material that can impact product yield. To compensate for such product loss, a continuous clarification of cell bleed material using AWS can be implemented at a steady-state condition during a long-term perfusion process17. This strategy may reduce the product loss in the bleed material and utilize more of the protein produced. In addition, cells after the continuous clarification using AWS could potentially be added back into the bioreactor if desired for higher cell density and productivity. Thus, continuous clarification of cell bleed material with AWS may offer an opportunity to increase yield and/or decrease secondary filter surface area in a given process.
In summary, the utility of AWS is not limited to simple primary clarification but may have utility for applications in continuous bioprocessing that may improve manufacturing rate and operational flexibility.

A critical step in a biomanufacturing process involving secreted therapeutic proteins is the removal of biomass from harvested cell culture fluid (HCCF). Traditionally, biomanufacturers have adopted centrifugation followed by depth filtration as their primary clarification methods in the production of monoclonal antibodies1. However, centrifugation may lead to high shear stress on the cells, resulting in increased cellular debris in the HCCF. This can potentially lead to filter fouling during filtration and result in extra contaminants postfiltration that can subsequently reduce downstream chromatography efficiency1,2,3. Furthermore, the customization of the centrifuges for a certain process can be costly and may require additional connections to clean-in-place and sterilize-in-place systems that may also be a limiting factor for scalability. The use of depth filters can compensate for the limitations of the centrifugation and also take advantage of single-use technology4. However, depth filters are primarily used as secondary clarification because they cannot withstand high cell culture densities5. Alternatively, tangential flow filtration (TFF) cell retention devices have been employed to mitigate shear stress but may experience challenges such as membrane polarization and poor harvest yields6. The above issues arising from the use of centrifugation plus depth filtration or TFF creates an opportunity for the improvement of the primary clarification process of HCCF.
Acoustic separation was introduced as a technology that can be utilized for harvesting secreted proteins from cell cultures with high-quality protein products7,8. Acoustic separation is achieved through the propagation and reflection of multidimensional standing waves that interact with suspended fluids and retained particles9,10. These particles experience three forces: fluid drag, gravity, and acoustic radiation. When each of the forces are equally opposed to each other, reaching equilibrium, the particles are suspended and trapped within the ultrasonic standing wave10. In a cell culture suspension, cells are held within this pressure node plane of standing waves, the node grows as cells coalesce, and eventually these clusters of cellular nodes fall from the gravitational force9. These sedimented cells are then removed from the media, which allows for the clarified media to be pumped out for further downstream processing. The utilization of ultrasonic waves as a separation method has started to translate into biological applications ranging from separation of lipid particles and red blood cells11 to mammalian perfusion cell culture12. With its relative capabilities to reduce costs, labor, and cellular stress by avoiding centrifugation, depth filtration, or TFF, biomanufacturers are exploring the potential applications of using acoustic separation.
This study provides a general protocol for operating a benchtop acoustic wave separator (AWS) for clarification of CHO cell culture, presents representative data, and demonstrates how to achieve effective cell clarification and product recovery.

<table><tbody><tr><td>Acoustophorectic chamber</td><td>Pall</td><td>CAS-AC-K1 Cadence acoustic chamber kit</td><td></td></tr><tr><td>ActiCHO P</td><td>GE</td><td>SH31025.01</td><td>powder medium</td></tr><tr><td>AWS</td><td>Pall</td><td>CAS-SYS (60500101-SP)</td><td></td></tr><tr><td>Cadence Acoustic Separator Software</td><td>Pall</td><td>Cadence Acoustic Separator Interface Ver. 1.0.4</td><td></td></tr><tr><td>CHO-K1 cells</td><td>VRC</td><td>VRC01</td><td></td></tr><tr><td>Computer</td><td>Dell</td><td>Latitude 3470</td><td>Windows 7, 64 bit OS</td></tr><tr><td>Isopropanol (70%)</td><td>LabChem</td><td>LC157605</td><td>20L prepped 70% IPA</td></tr><tr><td>L-glutamine</td><td>Corning</td><td>25-005-CV</td><td>200 mM stock solution</td></tr><tr><td>Masterflex L/S 14 tubing</td><td>Cole-Parmer</td><td>96400-14</td><td>peroxide-cured silicone tubing, 25ft</td></tr><tr><td>Masterflex L/S 16 tubing</td><td>Cole-Parmer</td><td>96400-16</td><td>peroxide-cured silicone tubing, 25ft</td></tr><tr><td>Tubing set</td><td>Pall</td><td>CAS-FP-K1 Tubing set</td><td></td></tr><tr><td>Turbidity probe</td><td>Pall</td><td>CAS-TS-S1 (60500106)</td><td></td></tr></tbody></table>

primary clarification of cho harvested cell culture fluid using an acoustic separator

Primary clarification is an essential step in a biomanufacturing process for the initial removal of cells from therapeutic products within the harvested cell culture fluid. While traditional methods like centrifugation or filtration are widely implemented for cell removal, the equipment for these processes have large footprints and operation can involve contamination risks and filter fouling. Additionally, traditional methods may not be ideal for continuous bioprocessing schemes for primary clarification. Thus, an alternate application using acoustic (sound) waves was investigated to continuously separate cells from the cell culture fluid. Presented in this study is a detailed protocol for using a bench-scale acoustic wave separator (AWS) for the primary separation of culture fluid containing a monoclonal IgG1 antibody from a CHO cell bioreactor harvest. Representative data are presented from the AWS and demonstrate how to achieve effective cell clarification and product recovery. Finally, potential applications for AWS in continuous bioprocessing are discussed. Overall, this study provides a practical and general protocol for the implementation of AWS in primary clarification for CHO cell cultures and further describes its application potential in continuous bioprocessing.

As described in the protocol, the AWS was used to clarify HCCF at a density of 12.4 x 106 cells/mL, as shown in Figure 1. The feed pump was set to 3.5 mL/min (5 L/day), representing a cell bleed rate within the presumed suitable range for a 10&#8211;20 L culture. As the HCCF entered the AWS chamber, the turbidity measurements from the feed turbidity probe remained consistent, around 1,000&#8211;1,100 NTU, and measurements from the probe1 turbidity probe remained around 40&#8211;50 NTU (Figure 8). Using the two measurements, cell removal efficiency
<img alt="Equation 2" src="/files/ftp_upload/61161/61161equ02.jpg" />
was calculated and averaged 95%. It was found that a turbidity measurement of 40&#8211;50 NTU was the minimum turbidity level achievable and thus no further separation from an additional AWS chamber in series was feasible.
While the AWS could separate with high efficiency at lower flow rates, keeping the HCCF for a longer time within the chamber caused temperature increases, which should be a consideration when selecting lower flow rates. Figure 9 is an example of the temperature difference of the HCCF before entering the acoustophoretic chamber and after acoustic separation at a feed flow rate of 3.5 mL/min, which showed a &#62;6 &#176;C increase in temperature due to prolonged time within the acoustic chamber.
Another important consideration when running high cell density harvests (i.e., &#62;20 x 106 cells/mL) is the saturation of the turbidity probes. The turbidity measurements for the feed turbidity probe became saturated over 4,400 NTU (Figure 10), which may result in underestimation in the calculation of cell removal efficiency.
To test the effect of different feed rates on the cell clarification, cell removal efficiency was measured at different feed rates. As shown in Figure 11, cell removal efficiency decreased significantly from ~100% to 57% as the feed pump rate increased. In general, the slower the feed pump rate, the better the cell clarification. However, optimization of feed rate is recommended for each application.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61161/61161fig01.jpg" /> 
Figure 1: AWS Setup.&#160;Once the pumps were turned on with the software (a), the HCCF was channeled in via a feed pump (b) through the feed turbidity probe (c) then to the AWS chamber (d). Inside the chamber, acoustic forces trapped cells from the flow in nodes of waves and caused clumping. Decreased buoyancy caused cells to drop through gravitational force, and cells were removed from the waste port of the acoustophoretic chamber via stage1 pump (e) to a cell harvest bottle (f) while the clarified material exited to the probe1 turbidity probe (c) through the permeate port of the chamber to a product harvest bottle (g). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61161/61161fig02.jpg" /> 
Figure 2: Back of AWS system.&#160;The turbidity probes and the chambers are connected to their respective ports at the back of AWS system via turbidity probe ethernet (a) and chamber power BNC (b) cables. Also, the computer is connected via PC ethernet cable (c). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61161/61161fig03.jpg" /> 
Figure 3: Turbidity probes and housing.&#160;Each turbidity probe (a) must be properly inserted to the respective turbidity meter and thermometer housing (b) and tightened with the screws. The chamber power BNC cable (c) should be connected to the back of the acoustophoretic chamber (d) only after the piezo transducer in the chamber is filled with fluid. The probes are indicated as following: F = feed turbidity, 1 = probe1 turbidity, and 2, 3, and 4 are unused probes (or can be used to serialize the procedure). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61161/61161fig04.jpg" /> 
Figure 4: Connection between the turbidity housing and the acoustophoretic chamber.&#160;The feed tubing is connected to the input of feed turbidity port (a) via the feed pump. The output of the feed turbidity port (b) is connected to the inlet ports of the acoustophoretic chamber (c) via y-tubing. The stage1 tubing is connected from the waste port of the acoustophoretic chamber (d) via the stage1 pump to a cell collection vessel. The permeate port of the acoustophoretic chamber (e) is connected to the input of probe1 turbidity port (f). The harvest tubing is connected from out of the probe1 turbidity port (g) to a product collection vessel. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61161/61161fig05.jpg" /> 
Figure 5: Readings panel in the Acoustic Separator software.&#160;The program has two panels, &#8220;Readings&#8221; and &#8220;Controls&#8221;. Within the &#8220;Readings&#8221; panel, turbidity (a), temperature (b), and percent reduction (c) are monitored. To initiate the data recording, the &#8220;Start Test&#8221; button (d) needs to be clicked, which will change the button&#8217;s color to green. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61161/61161fig06.jpg" /> 
Figure 6: Controls panel in the program.&#160;Within the &#8220;Controls&#8221; panel, the pumps can be turned on or off, and the rate of pumping can be changed for feed (a) and other stages (b). Also, the chamber power on the piezo transducer can be turned on or off (c) and be changed with a slide bar (d). The experiments used 10 W, because it is the recommended power setting for CHO cells as recommended by the manufacturer. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61161/61161fig07.jpg" /> 
Figure 7: AWS Chamber.&#160;Once the cells were inside the chamber, the acoustic forces trapped cells in nodes of waves and caused cells to cluster (a). These cell clusters increased in size until they lost their buoyancy and eventually settled down by gravitational force (b). Next, the settled cells were removed from the waste port of the acoustophoretic chamber via stage1 pump to a cell harvest bottle (c). The product exited the chamber through permeate port (d) while HCCF continuously filled the chamber via inlet ports (e). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/61161/61161fig08.jpg" /> 
Figure 8: Turbidity measurements for a 12 x 106 cell/mL CHO cell culture with a feed pump rate of 3.5 mL/min.&#160;Feed turbidity (blue) was approximately 1,000 NTU and permeate exiting the stage 1 turbidity meter (orange) was 40&#8211;60 NTU. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/61161/61161fig09.jpg" /> 
Figure 9: Temperature measurements for a 12 x 106 cell/mL CHO cell culture with a feed pump rate of 3.5 mL/min.&#160;Feed temperature (blue) was approximately 21 &#176;C and permeate exiting the stage 1 turbidity meter (orange) was approximately 27 &#176;C. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/61161/61161fig10.jpg" /> 
Figure 10: Turbidity measurement example for a high cell density sample.&#160;When the cell density was &#62;20 x 106 cells/mL, the feed turbidity measurement was saturated at the maximum value of 4,400 NTU, resulting in underestimation of cell removal efficiency. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/61161/61161fig11.jpg" /> 
Figure 11: Cell removal efficiency comparison.&#160;As the feed rate increased, the cell separation decreased. Hence, as the feed rate increased, the cell clarification efficiency decreased. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61161/61161fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Parameter</td>
			<td>Specification</td>
		</tr>
		<tr>
			<td>Flow Rate</td>
			<td>0 &#8211; 10 L/h</td>
		</tr>
		<tr>
			<td>Pressure Range</td>
			<td>0 &#8211; 2 bar (0 &#8211; 30 psi)</td>
		</tr>
		<tr>
			<td>Feed Fluid Temperature</td>
			<td>0 &#8211; 40 &#176;C (32 &#8211; 104 &#176;F)</td>
		</tr>
		<tr>
			<td>Operating Temperature</td>
			<td>0 &#8211; 40 &#176;C (32 &#8211; 104 &#176;F)</td>
		</tr>
	</tbody>
</table>
Table 1: Operating Conditions. The recommended operating conditions from the AWS manufacturer for flow rate, pressure range, feed fluid, and operating temperature.

Watch this Scientific Journal Video about Primary Clarification of CHO Harvested Cell Culture Fluid using an Acoustic Separator at JoVE.com

Primary Clarification of CHO Harvested Cell Culture Fluid using an Acoustic Separator

Presented here is a protocol for the primary clarification of CHO cell culture using an acoustic separator. This protocol can be used for the primary clarification of shake flask cultures or bioreactor harvests and has the potential application for continuous clarification of the cell bleed material during perfusion bioreactor operations.

Primary clarification is an essential step in a biomanufacturing process for the initial removal of cells from therapeutic products within the harvested ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

5.0K Views.  U.S. Food and Drug Administration. Presented here is a protocol for the primary clarification of CHO cell culture using an acoustic separator. This protocol can be used for the primary clarification of shake flask cultures or bioreactor harvests and has the potential application for continuous clarification of the cell bleed material during perfusion bioreactor operations.

Video: Primary Clarification of CHO Harvested Cell Culture Fluid using an Acoustic Separator

Preparation of Primary Acute Lymphoblastic Leukemia Cells in Different Cell Cycle Phases by Centrifugal Elutriation

The ability to synchronize cells has been central to advancing our understanding of cell cycle regulation. Common techniques employed include serum deprivation; chemicals which arrest cells at different cell cycle phases; or the use of mitotic shake-off which exploits their reduced adherence. However, all of these have disadvantages. For example, serum starvation works well for normal cells but less well for tumor cells with compromised cell cycle checkpoints due to oncogene activation or tumor suppressor loss. Similarly, chemically-treated cell populations can harbor drug-induced damage and show stress-related alterations. A technique which circumvents these problems is counterflow centrifugal elutriation (CCE), where cells are subjected to two opposing forces, centrifugal force and fluid velocity, which results in the separation of cells on the basis of size and density. Since cells advancing through the cycle typically enlarge, CCE can be used to separate cells into different cell cycle phases. Here we apply this technique to primary acute lymphoblastic leukemia cells. Under optimal conditions, an essentially pure population of cells in G1 phase and a highly enriched population of cells in G2/M phases can be obtained in excellent yield. These cell populations are ideally suited for studying cell cycle-dependent mechanisms of action of anticancer drugs and for other applications. We also show how modifications to the standard procedure can result in suboptimal performance and discuss the limitations of the technique. The detailed methodology presented should facilitate application and exploration of the technique to other types of cells.

This protocol describes the use of centrifugal elutriation to separate primary acute lymphoblastic leukemia cells into different cell cycle phases.

The ability to synchronize cells has been central to advancing our understanding of cell cycle regulation. Common techniques employed include serum ...

Cancer Research

Isolation and Enrichment of Liver Progenitor Subsets Identified by a Novel Surface Marker Combination

During chronic liver injuries, progenitor cells expand in a process called ductular reaction, which also entails the appearance of inflammatory cellular infiltrate and epithelial cell activation. The progenitor cell population during such inflammatory reactions has mostly been investigated using single surface markers, either by histological analysis or by flow cytometry-based techniques. However, novel surface markers identified various functionally distinct subsets within the liver progenitor/stem cell compartment. The method presented here describes the isolation and detailed flow cytometry analysis of progenitor subsets using novel surface marker combinations. Moreover, it demonstrates how the various progenitor cell subsets can be isolated with high purity using automated magnetic and FACS sorting-based methods. Importantly, novel and simplified enzymatic dissociation of the liver allows for the isolation of these rare cell populations with a high viability that is superior in comparison to other existing methods. This is especially relevant for further studying progenitor cells in vitro or for isolating high-quality RNA to analyze the gene expression profile.

Liver injuries are accompanied by progenitor cell expansion that represents a heterogeneous cell population. Novel classification of this cellular compartment allows for the distinguishing of multiple subsets. The method described here illustrates the flow cytometry analysis and high purity isolation of various subsets that can be used for further assays.

During chronic liver injuries, progenitor cells expand in a process called ductular reaction, which also entails the appearance of inflammatory cellular ...

Developmental Biology

A Microfluidic Platform for Precision Small-volume Sample Processing and Its Use to Size Separate Biological Particles with an Acoustic Microdevice

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. However, to achieve robust sample manipulation it is necessary to address device integration with the macroscale environment. To realize repeatable, sensitive particle separation with microfluidic devices, this protocol presents a complete automated and integrated microfluidic platform that enables precise processing of 0.15&#8211;1.5 ml samples using microfluidic devices. Important aspects of this system include modular device layout and robust fixtures resulting in reliable and flexible world to chip connections, and fully-automated fluid handling which accomplishes closed-loop sample collection, system cleaning and priming steps to ensure repeatable operation. Different microfluidic devices can be used interchangeably with this architecture. Here we incorporate an acoustofluidic device, detail its characterization, performance optimization, and demonstrate its use for size-separation of biological samples. By using real-time feedback during separation experiments, sample collection is optimized to conserve and concentrate sample. Although requiring the integration of multiple pieces of equipment, advantages of this architecture include the ability to process unknown samples with no additional system optimization, ease of device replacement, and precise, robust sample processing.

This protocol describes a system architecture for performing automated small volume (0.15&#8211;1.5 ml) particle separations using a microfluidic device, and discusses methods to optimize acoustofluidic device performance and operation.

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. ...

A Microfluidic Platform for High-throughput Single-cell Isolation and Culture

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet high-throughput, method to perform single-cell culture experiments. Here, we report a microfluidic chip-based strategy for high-efficiency single-cell isolation (~77%) and demonstrate its capability of performing long-term single-cell culture (up to 7 d) and cellular heterogeneity analysis using clonogenic assay. These applications were demonstrated with KT98 mouse neural stem cells, and A549 and MDA-MB-435 human cancer cells. High single-cell isolation efficiency and long-term culture capability are achieved by using different sizes of microwells on the top and bottom of the microfluidic channel. The small microwell array is designed for precisely isolating single-cells, and the large microwell array is used for single-cell clonal culture in the microfluidic chip. This microfluidic platform constitutes an attractive approach for single-cell culture applications, due to its flexibility of adjustable cell culture spaces for different culture strategies, without decreasing isolation efficiency.

Here, we present a protocol for isolating and culturing single cells with a microfluidic platform, which utilizes a new microwell design concept to allow for high-efficiency single cell isolation and long-term clonal culture.

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet ...

Microfluidic Buffer Exchange for Interference-free Micro/Nanoparticle Cell Engineering

Engineering cells with active-ingredient-loaded micro/nanoparticles (NPs) is becoming an increasingly popular method to enhance native therapeutic properties, enable bio imaging and control cell phenotype. A critical yet inadequately addressed issue is the significant number of particles that remain unbound after cell labeling which cannot be readily removed by conventional centrifugation. This leads to an increase in bio imaging background noise and can impart transformative effects onto neighboring non-target cells. In this protocol, we present an inertial microfluidics-based buffer exchange strategy termed as Dean Flow Fractionation (DFF) to efficiently separate labeled cells from free NPs in a high throughput manner. The developed spiral microdevice facilitates continuous collection (&#62;90% cell recovery) of purified cells (THP-1 and MSCs) suspended in new buffer solution, while achieving &#62;95% depletion of unbound fluorescent dye or dye-loaded NPs (silica or PLGA). This single-step, size-based cell purification strategy enables high cell processing throughput (106 cells/min) and is highly useful for large-volume cell purification of micro/nanoparticle engineered cells to achieve interference-free clinical application.

This protocol describes the use of an inertial microfluidics-based buffer exchange strategy to purify micro/nanoparticle engineered cells with efficient depletion of unbound particles.

Engineering cells with active-ingredient-loaded micro/nanoparticles (NPs) is becoming an increasingly popular method to enhance native therapeutic ...

Microfluidic Acoustophoresis for Flowthrough Separation of Gram-Negative Bacteria using Aptamer Affinity Beads

This article describes the fabrication and operation of microfluidic acoustophoretic chips using a microfluidic acoustophoresis technique and aptamer-modified microbeads that can be used for the fast, efficient isolation of Gram-negative bacteria from a medium. This method enhances the separation efficiency using a mix of long, square microchannels. In this system, the sample and buffer are injected into the inlet port through a flow controller. For bead centering and sample separation, AC power is applied to the piezoelectric transducer via a function generator with a power amplifier to generate acoustic radiation force in the microchannel. There is a bifurcated channel at both the inlet and outlet, enabling simultaneous separation, purification, and concentration. The device has a recovery rate of &gt;98% and purity of 97.8% up to a 10x dose concentration. This study has demonstrated a recovery rate and purity higher than the existing methods for separating bacteria, suggesting that the device can separate bacteria efficiently.

This paper describesthe fabrication and operation of microfluidic acoustophoretic chips using the microfluidic acoustophoresis technique and aptamer-modified microbeads that can be used for fast, efficient isolation of Gram-negative bacteria from a medium.

This article describes the fabrication and operation of microfluidic acoustophoretic chips using a microfluidic acoustophoresis technique and ...

Engineering

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

Biology

Size Exclusion Chromatography for Separating Extracellular Vesicles from Conditioned Cell Culture Media

Extracellular vesicles (EVs) are nano-sized lipid-membrane bound structures that are released from all cells, are present in all biofluids, and contain proteins, nucleic acids, and lipids that are reflective of the parent cell from which they are derived. Proper separation of EVs from other components in a sample allows for characterization of their associated cargo and lends insight into their potential as intercellular communicators and non-invasive biomarkers for numerous diseases. In the current study, oligodendrocyte derived EVs were isolated from cell culture media using a combination of state-of-the-art techniques, including ultrafiltration and size exclusion chromatography (SEC) to separate EVs from other extracellular proteins and protein complexes. Using commercially available SEC columns, EVs were separated from extracellular proteins released from human oligodendroglioma cells under both control and endoplasmic reticulum (ER) stress conditions. The canonical EV markers CD9, CD63, and CD81 were observed in fractions 1-4, but not in fractions 5-8. GM130, a protein of the Golgi apparatus, and calnexin, an integral protein of the ER, were used as negative EV markers, and were not observed in any fraction. Further, when pooling and concentrating fractions 1-4 as the EV fraction, and fractions 5-8 as the protein fraction, expression of CD63, CD81, and CD9 in the EV fraction was observed. The expression of GM130 or calnexin was not observed in either of the fraction types. The pooled fractions from both control and ER stress conditions were visualized with transmission electron microscopy and vesicles were observed in the EV fractions, but not in the protein fractions. Particles in the EV and protein fractions from both conditions were also quantified with nanoparticle tracking analysis. Together, these data demonstrate that SEC is an effective method for separating EVs from conditioned cell culture media.

The protocol here demonstrates that extracellular vesicles can be adequately separated from conditioned cell culture media using size exclusion chromatography.

Extracellular vesicles (EVs) are nano-sized lipid-membrane bound structures that are released from all cells, are present in all biofluids, and contain ...

Neuroscience

Harvesting and Disaggregation: An Overlooked Step in Biofilm Methods Research

Biofilm methods consist of four distinct steps: growing the biofilm in a relevant model, treating the mature biofilm, harvesting the biofilm from the surface and disaggregating the clumps, and analyzing the sample. Of the four steps, harvesting and disaggregation are the least studied but nonetheless critical when considering the potential for test bias. This article demonstrates commonly used harvesting and disaggregation techniques for biofilm grown on three different surfaces. The three biofilm harvesting and disaggregation techniques, gleaned from an extensive literature review, include vortexing and sonication, scraping and homogenization, and scraping, vortexing and sonication. Two surface types are considered: hard non-porous (polycarbonate and borosilicate glass) and porous (silicone). Additionally, we provide recommendations for the minimum information that should be included when reporting the harvesting technique followed and an accompanying method to check for bias.

This paper details methods that demonstrate three common biofilm harvesting and disaggregation techniques on two surface types, ruggedness testing of a harvesting method and minimum information to consider when choosing and optimizing harvesting and disaggregation techniques to increase reproducibility.

Biofilm methods consist of four distinct steps: growing the biofilm in a relevant model, treating the mature biofilm, harvesting the biofilm from the ...

Leukemic Subpopulation Harvest: A Method for Spatial Separation of Leukemic Cell Subpopulations from 2D Co-culture

Source: Slone, W. L., et al. <a href="https://jove.unicartagenaproxy.elogim.com/t/53645">Modeling Chemotherapy Resistant Leukemia In Vitro.</a> J. Vis. Exp. (2016).
This video describes a method for spatial separation of three leukemic cell subpopulations: the suspended leukemic cells freely floating in the media, phase bright leukemic cells adhered to the surface of the mesenchymal stromal cell monolayer, and the phase dim leukemic cells that have migrated beneath the mesenchymal stromal cell monolayer from 2D co-culture.

Source: Slone, W. L., et al. Modeling Chemotherapy Resistant Leukemia In Vitro. J. Vis. Exp. (2016).
This video describes a method for spatial separation ...

Primary Clarification of CHO Harvested Cell Culture Fluid using an Acoustic Separator

Summary

Explore More Videos

Chapters in this video

Preparation of Primary Acute Lymphoblastic Leukemia Cells in Different Cell Cycle Phases by Centrifugal Elutriation

Isolation and Enrichment of Liver Progenitor Subsets Identified by a Novel Surface Marker Combination

A Microfluidic Platform for Precision Small-volume Sample Processing and Its Use to Size Separate Biological Particles with an Acoustic Microdevice

A Microfluidic Platform for High-throughput Single-cell Isolation and Culture

Microfluidic Buffer Exchange for Interference-free Micro/Nanoparticle Cell Engineering

Microfluidic Acoustophoresis for Flowthrough Separation of Gram-Negative Bacteria using Aptamer Affinity Beads

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Size Exclusion Chromatography for Separating Extracellular Vesicles from Conditioned Cell Culture Media

Harvesting and Disaggregation: An Overlooked Step in Biofilm Methods Research

Leukemic Subpopulation Harvest: A Method for Spatial Separation of Leukemic Cell Subpopulations from 2D Co-culture