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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A., Suda, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Harvest and recovery of monoclonal antibodies: cell removal and clarification. 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Z., Lanthier, S., Kamen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acoustic+cell+filter:+a+proven+cell+retention+technology+for+perfusion+of+animal+cell+cultures.">Acoustic cell filter: a proven cell retention technology for perfusion of animal cell cultures.</a> Biotechnology Advances. 22 (6), 433-444 (2004).</li><li>Wolf, M. K. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+Performance+in+Mammalian+Cell+Perfusion+Cultures+by+Growth+Inhibition.">Improved Performance in Mammalian Cell Perfusion Cultures by Growth Inhibition.</a> Biotechnology Journal. 14 (2), 1700722 (2019).</li><li>Lin, H., Leighty, R. W., Godfrey, S., Wang, S. 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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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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://www-jove-com.remotexs.ntu.edu.sg/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 ...

primary-clarification-cho-harvested-cell-culture-fluid-using-an

Nanyang Technological University

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

Developing Custom Chinese Hamster Ovary-host Cell Protein Assays using Acoustic Membrane Microparticle Technology

Custom assays for unique proteins are often limited to time consuming manual detection and quantitation techniques such as ELISA or Western blots due to the complexity of development on alternate platforms. BioScale's proprietary Acoustic Membrane MicroParticle (AMMP) technology allows sandwich immunoassays to be easily developed for use on the ViBE platform, providing better sensitivity, reproducibility, and automated operation. Provided as an example, this protocol outlines the procedure for developing a custom Chinese Hamster Ovary- Host Cell Protein (CHO-HCP) assay. The general principles outlined here can be followed for the development of a wide variety of immunoassays.
An AMMP assay measures antigen concentration by measuring changes in oscillation frequency caused by the binding of microparticles to the sensor surface to calculate. It consists of four major components: (1) a cartridge that contains a functionalized eight sensor chip (2) antibody labeled magnetic microparticles, (3) hapten tagged antibody that binds to the surface of the functionalized chip (4) samples containing the antigen of interest. BioScale's biosensor is a resonant device that contains eight individual membranes with separate fluidic paths. The membranes change oscillation frequency in response to mass accumulating on the surface and this frequency change is used to quantitate the amount of added mass.
To facilitate use in a wide variety of immunoassays the sensor is functionalized with an anti-hapten antibody. Assay specific antibodies are modified through the covalent conjugation of a hapten tag to one antibody and biotin to the other. The biotin label is used to bind the antibody to streptavidin coupled magnetic beads which, in combination with the hapten-tagged antibody, are used to capture the analyte in a sandwich. The complex binds to the chip through the anti-hapten/hapten interaction. At the end of each assay run the sensors are cleaned with a dilute acid enabling the sequential analysis of columns from a 96-well plate.
Here, we present the method for developing a custom CHO-HCP AMMP assay for bioprocess development. Developing AMMP assays or modifying existing assays into AMMP assays can provide better performance (reproducibility, sensitivity) in complex samples and reduced operator time. The protocol shows the steps for development and the discussion section reviews representative results. For a more in-depth explanation of assay optimization and customization parameters contact BioScale. This kit offers generic bioprocess development assays such as Residual Protein A, Product titer, and CHO-HCP.

Development of custom assays on the ViBE platform for more sensitive, reproducible, automated results in complex matrices is described. The universal cartridge allows assays to be easily adapted for use with custom assays. This versatility enables rapid development and validation of novel assays or automated versions of existing manual assays, exemplified in this video.

Custom assays for unique proteins are often limited to time consuming manual detection and quantitation techniques such as ELISA or Western blots due to ...

Increasing cDNA Yields from Single-cell Quantities of mRNA in Standard Laboratory Reverse Transcriptase Reactions using Acoustic Microstreaming

Correlating gene expression with cell behavior is ideally done at the single-cell level. However, this is not easily achieved because the small amount of labile mRNA present in a single cell (1-5% of 1-50pg total RNA, or 0.01-2.5pg mRNA, per cell 1) mostly degrades before it can be reverse transcribed into a stable cDNA copy. For example, using standard laboratory reagents and hardware, only a small number of genes can be qualitatively assessed per cell 2. One way to increase the efficiency of standard laboratory reverse transcriptase (RT) reactions (i.e. standard reagents in microliter volumes) comprising single-cell amounts of mRNA would be to more rapidly mix the reagents so the mRNA can be converted to cDNA before it degrades. However this is not trivial because at microliter scales liquid flow is laminar, i.e. currently available methods of mixing (i.e. shaking, vortexing and trituration) fail to produce sufficient chaotic motion to effectively mix reagents. To solve this problem, micro-scale mixing techniques have to be used 3,4. A number of microfluidic-based mixing technologies have been developed which successfully increase RT reaction yields 5-8. However, microfluidics technologies require specialized hardware that is relatively expensive and not yet widely available. A cheaper, more convenient solution is desirable. The main objective of this study is to demonstrate how application of a novel "micromixing" technique to standard laboratory RT reactions comprising single-cell quantities of mRNA significantly increases their cDNA yields. We find cDNA yields increase by approximately 10-100-fold, which enables: (1) greater numbers of genes to be analyzed per cell; (2) more quantitative analysis of gene expression; and (3) better detection of low-abundance genes in single cells. The micromixing is based on acoustic microstreaming 9-12, a phenomenon where sound waves propagating around a small obstacle create a mean flow near the obstacle. We have developed an acoustic microstreaming-based device ("micromixer") with a key simplification; acoustic microstreaming can be achieved at audio frequencies by ensuring the system has a liquid-air interface with a small radius of curvature 13. The meniscus of a microliter volume of solution in a tube provides an appropriately small radius of curvature. The use of audio frequencies means that the hardware can be inexpensive and versatile 13, and nucleic acids and other biochemical reagents are not damaged like they can be with standard laboratory sonicators.

We describe a novel method for increasing cDNA yield from single-cell quantities of mRNA in otherwise standard laboratory reverse transcription reactions. The novelty resides in the use of a micromixer, which utilizes the phenomenon of acoustic microstreaming, to mix fluids at microliter scales more effectively than shaking, vortexing or trituration.

Correlating gene expression with cell behavior is ideally done at the single-cell level.  However, this is not easily achieved because the small amount of ...

Three-dimensional Cell Culture Model for Measuring the Effects of Interstitial Fluid Flow on Tumor Cell Invasion

The growth and progression of most solid tumors depend on the initial transformation of the cancer cells and their response to stroma-associated signaling in the tumor microenvironment 1. Previously, research on the tumor microenvironment has focused primarily on tumor-stromal interactions 1-2. However, the tumor microenvironment also includes a variety of biophysical forces, whose effects remain poorly understood. These forces are biomechanical consequences of tumor growth that lead to changes in gene expression, cell division, differentiation and invasion3. Matrix density 4, stiffness 5-6, and structure 6-7, interstitial fluid pressure 8, and interstitial fluid flow 8 are all altered during cancer progression.
Interstitial fluid flow in particular is higher in tumors compared to normal tissues 8-10. The estimated interstitial fluid flow velocities were measured and found to be in the range of 0.1-3 &mu;m s-1, depending on tumor size and differentiation 9, 11. This is due to elevated interstitial fluid pressure caused by tumor-induced angiogenesis and increased vascular permeability 12. Interstitial fluid flow has been shown to increase invasion of cancer cells 13-14, vascular fibroblasts and smooth muscle cells 15. This invasion may be due to autologous chemotactic gradients created around cells in 3-D 16 or increased matrix metalloproteinase (MMP) expression 15, chemokine secretion and cell adhesion molecule expression 17. However, the mechanism by which cells sense fluid flow is not well understood. In addition to altering tumor cell behavior, interstitial fluid flow modulates the activity of other cells in the tumor microenvironment. It is associated with (a) driving differentiation of fibroblasts into tumor-promoting myofibroblasts 18, (b) transporting of antigens and other soluble factors to lymph nodes 19, and (c) modulating lymphatic endothelial cell morphogenesis 20.
The technique presented here imposes interstitial fluid flow on cells in vitro and quantifies its effects on invasion (Figure 1). This method has been published in multiple studies to measure the effects of fluid flow on stromal and cancer cell invasion 13-15, 17. By changing the matrix composition, cell type, and cell concentration, this method can be applied to other diseases and physiological systems to study the effects of interstitial flow on cellular processes such as invasion, differentiation, proliferation, and gene expression.

Interstitial fluid flow is elevated in solid tumors and can modulate tumor cell invasion. Here we describe a technique to apply interstitial fluid flow to cells embedded in a matrix and then measure its effects on cell invasion. This technique can be easily adapted to study other systems.

The growth and progression of most solid tumors depend on the initial transformation of the cancer cells and their response to stroma-associated signaling ...

Cell Co-culture Patterning Using Aqueous Two-phase Systems

Cell patterning technologies that are fast, easy to use and affordable will be required for the future development of high throughput cell assays, platforms for studying cell-cell interactions and tissue engineered systems. This detailed protocol describes a method for generating co-cultures of cells using biocompatible solutions of dextran (DEX) and polyethylene glycol (PEG) that phase-separate when combined above threshold concentrations. Cells can be patterned in a variety of configurations using this method. Cell exclusion patterning can be performed by printing droplets of DEX on a substrate and covering them with a solution of PEG containing cells. The interfacial tension formed between the two polymer solutions causes cells to fall around the outside of the DEX droplet and form a circular clearing that can be used for migration assays. Cell islands can be patterned by dispensing a cell-rich DEX phase into a PEG solution or by covering the DEX droplet with a solution of PEG. Co-cultures can be formed directly by combining cell exclusion with DEX island patterning. These methods are compatible with a variety of liquid handling approaches, including manual micropipetting, and can be used with virtually any adherent cell type.

Aqueous two-phase systems were used to simultaneously pattern multiple populations of cells. This fast and easy method for cell patterning takes advantage of the phase separation of aqueous solutions of dextran and polyethylene glycol and the interfacial tension that exists between the two polymer solutions.

Cell patterning  technologies that are fast, easy to use and affordable will be required for the  future development of high throughput cell assays, ...

Cell Squeezing as a Robust, Microfluidic Intracellular Delivery Platform

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This technology has shown potential in addressing previously challenging applications; including, delivery to primary immune cells, cell reprogramming, carbon nanotube, and quantum dot delivery. This vector-free microfluidic platform relies on mechanical disruption of the cell membrane to facilitate cytosolic delivery of the target material. Herein, we describe the detailed method of use for these microfluidic devices including, device assembly, cell preparation, and system operation. This delivery approach requires a brief optimization of device type and operating conditions for previously unreported applications. The provided instructions are generalizable to most cell types and delivery materials as this system does not require specialized buffers or chemical modification/conjugation steps. This work also provides recommendations on how to improve device performance and trouble-shoot potential issues related to clogging, low delivery efficiencies, and cell viability.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This protocol provides detailed steps on how to use the system for a broad range of applications.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This ...

Microfluidic Flow Chambers Using Reconstituted Blood to Model Hemostasis and Platelet Transfusion In Vitro

Blood platelets prepared for transfusion gradually lose hemostatic function during storage. Platelet function can be investigated using a variety of (indirect) in vitro experiments, but none of these is as comprehensive as microfluidic flow chambers. In this protocol, the reconstitution of thrombocytopenic fresh blood with stored blood bank platelets is used to simulate platelet transfusion. Next, the reconstituted sample is perfused in microfluidic flow chambers which mimic hemostasis on exposed subendothelial matrix proteins. Effects of blood donation, transport, component separation, storage and pathogen inactivation can be measured in paired experimental designs. This allows reliable comparison of the impact every manipulation in blood component preparation has on hemostasis. Our results demonstrate the impact of temperature cycling, shear rates, platelet concentration and storage duration on platelet function. In conclusion, this protocol analyzes the function of blood bank platelets and this ultimately aids in optimization of the processing chain including phlebotomy, transport, component preparation, storage and transfusion.

Microfluidic Flow Chambers Using Reconstituted Blood to Model Hemostasis and Platelet Transfusion In Vitro

Platelet transfusion and hemostasis was modeled using blood reconstitution and microfluidic flow chambers to investigate the function of blood banking platelets. The data demonstrate the consequences of platelet storage lesion on hemostasis, in vitro.

Blood platelets prepared for transfusion gradually lose hemostatic function during storage. Platelet function can be investigated using a variety of ...

Fabrication of Inverted Colloidal Crystal Poly(ethylene glycol) Scaffold: A Three-dimensional Cell Culture Platform for Liver Tissue Engineering

The ability to maintain hepatocyte function in vitro, for the purpose of testing xenobiotics&#39; cytotoxicity, studying virus infection and developing drugs targeted at the liver, requires a platform in which cells receive proper biochemical and mechanical cues. Recent liver tissue engineering systems have employed three-dimensional (3D) scaffolds composed of synthetic or natural hydrogels, given their high water retention and their ability to provide the mechanical stimuli needed by the cells. There has been growing interest in the inverted colloidal crystal (ICC) scaffold, a recent development, which allows high spatial organization, homotypic and heterotypic cell interaction, as well as cell-extracellular matrix (ECM) interaction. Herein, we describe a protocol to fabricate the ICC scaffold using poly (ethylene glycol) diacrylate (PEGDA) and the particle leaching method. Briefly, a lattice is made from microsphere particles, after which a pre-polymer solution is added, properly polymerized, and the particles are then removed, or leached, using an organic solvent (e.g., tetrahydrofuran). The dissolution of the lattice results in a highly porous scaffold with controlled pore sizes and interconnectivities that allow media to reach cells more easily. This unique structure allows high surface area for the cells to adhere to as well as easy communication between pores, and the ability to coat the PEGDA ICC scaffold with proteins also shows a marked effect on cell performance. We analyze the morphology of the scaffold as well as the hepatocarcinoma cell (Huh-7.5) behavior in terms of viability and function to explore the effect of ICC structure and ECM coatings. Overall, this paper provides a detailed protocol of an emerging scaffold that has wide applications in tissue engineering, especially liver tissue engineering.

Fabrication of Inverted Colloidal Crystal Polyethylene glycol Scaffold: A Three-dimensional Cell Culture Platform for Liver Tissue Engineering

This manuscript presents a detailed protocol for the fabrication of an emerging three-dimensional hepatocyte culture platform, the inverted colloidal crystal scaffold, and the concomitant techniques to assess hepatocyte behavior. The size-controllable pores, interconnectivity and ability to conjugate extracellular matrix proteins to the poly(ethylene glycol) (PEG) scaffold enhance Huh-7.5 cell performance.

The ability to maintain hepatocyte function in vitro, for the purpose of testing xenobiotics&#39; cytotoxicity, studying virus infection and developing ...

Fabrication and Validation of an Organ-on-chip System with Integrated Electrodes to Directly Quantify Transendothelial Electrical Resistance

Organs-on-chips, in vitro models involving the culture of (human) tissues inside microfluidic devices, are rapidly emerging and promise to provide useful research tools for studying human health and disease. To characterize the barrier function of cell layers cultured inside organ-on-chip devices, often transendothelial or transepithelial electrical resistance (TEER) is measured. To this end, electrodes are usually integrated into the chip by micromachining methods to provide more stable measurements than is achieved with manual insertion of electrodes into the inlets of the chip. However, these electrodes frequently hamper visual inspection of the studied cell layer or require expensive cleanroom processes for fabrication. To overcome these limitations, the device described here contains four easily integrated electrodes that are placed and fixed outside of the culture area, making visual inspection possible. Using these four electrodes the resistance of six measurement paths can be quantified, from which the TEER can be directly isolated, independent of the resistance of culture medium-filled microchannels. The blood-brain barrier was replicated in this device and its TEER was monitored to show the device applicability. This chip, the integrated electrodes and the TEER determination method are generally applicable in organs-on-chips, both to mimic other organs or to be incorporated into existing organ-on-chip systems.

This publication describes the fabrication of an organ-on-chip device with integrated electrodes for direct quantification of transendothelial electrical resistance (TEER). For validation, the blood-brain barrier was mimicked inside this microfluidic device and its barrier function was monitored. The presented methods for electrode integration and direct TEER quantification are generally applicable.

Organs-on-chips, in vitro models involving the culture of (human) tissues inside microfluidic devices, are rapidly emerging and promise to provide useful ...

Handling and Assessment of Human Primary Prostate Organoid Culture

This paper describes a detailed protocol for three-dimensional (3D) culturing, handling, and evaluation of human primary prostate organoids. The process involves seeding of epithelial cells sparsely in a 3D matrix gel on a 96-well microplate with media changes to cultivate expansion into organoids. Morphology is then assessed by whole-well capturing of z-stack images. Compression of z-stacks creates a single in-focus image from which organoids are measured to quantify a variety of outputs, including circularity, roundness, and area.DNA, RNA, and protein can be collected from organoids recovered from the matrix gel. Cell populations of interest can be assessed by organoid dissociation and flow cytometry. Formalin-fixation-paraffin-embedding (FFPE) followed by sectioning is used for the histological assessment and antibody staining. Whole-mount immunofluorescent staining preserves organoid morphology and facilitates observation of protein localization in organoids in situ. Commercial assays that are traditionally used for 2D monolayer cells can be modified for 3D organoids. Used together, the techniques in this protocol provide a robust toolbox to quantify prostate organoid growth, morphologic characteristics, and expression of differentiation markers.

Here, we present a protocol to guide human primary prostate organoid handling then suggest endpoints to assess phenotype. Seeding, culture maintenance, recovery from matrix gel, morphologic quantification, embedding and sectioning, FFPE sectioning, whole-mount staining, and application of commercial assays are described.

This paper describes a detailed protocol for three-dimensional (3D) culturing, handling, and evaluation of human primary prostate organoids. The process ...

Screening and Identification of Small Peptides Targeting Fibroblast Growth Factor Receptor2 using a Phage Display Peptide Library

The human Fibroblast Growth Factor Receptor (FGFR) family comprises four members, namely, FGFR1, FGFR2, FGFR3, and FGFR4, that are involved in various biological activities including cell proliferation, survival, migration and differentiation. Several aberrations in the FGFR signaling pathway, owing to mutations or gene amplification events, have been identified in various types of cancers. Hence, recent research has focused on developing strategies involving therapeutic targeting of FGFRs. Current FGFR inhibitors at various stages of pre-clinical and clinical development include either small molecule inhibitors of tyrosine kinases or monoclonal antibodies, with only a few peptide- based inhibitors in the pipeline. Here, we provide a protocol using phage display technology to screen small peptides as antagonists of FGFR2. Briefly, a library of phage-displayed peptides was incubated in a plate coated with FGFR2. Subsequently, unbound phage was washed off by TBST (TBS + 0.1% [v/v] Tween-20), and bound phage was eluted with 0.2 M glycine-HCl buffer (pH 2.2). The eluted phage was further amplified and used as input for the next round of biopanning. Following three rounds of biopanning, the peptide sequences of individual phage clones were identified by DNA sequencing. Finally, the screened peptides were synthesized and analyzed for affinity and biological activity.

Herein, we present a detailed protocol for screening small peptides that bind to FGFR2 using a phage display peptide library. We further analyze the affinity of the selected peptides toward FGFR2 in vitro and its ability to suppress cell proliferation.

The human Fibroblast Growth Factor Receptor (FGFR) family comprises four members, namely, FGFR1, FGFR2, FGFR3, and FGFR4, that are involved in various ...