Name: operation of a benchtop bioreactor
Uploaded: 2013-09-12T15:45:00
Duration: 12 min 54 s
Description: Watch this Scientific Journal Video about Operation of a Benchtop Bioreactor at JoVE.com

This project was support in part by Johns Hopkins University, Office of the Provost through the Gateway Science Initiative.

<ol>
	<li>Begin operation with vessel setup. A conventional fermentor has the following components that need to be installed (Figure 1):
	<ol>
		<li>pH probe - for measurement of pH in the live culture. Calibrate probe before installation and sterilize with the vessel. Install probe in the headplate.</li>
		<li>pO2 probe - for measurement of dissolved oxygen content inside the vessel during fermentation. Install in the headplate.</li>
		<li>Harvest pipe for sampling the culture. Install this adjustable height component in the headplate.</li>
		<li>Gas sparger - resides at the bottom of the vessel and provides gas infusion to the culture. Hook sparger up to the gas source on the fermentor and mount in the headplate.</li>
		<li>Impeller shaft and impeller - the impeller stirs the culture and is critical for maintaining culture uniformity in the vessel.</li>
		<li>Exhaust gas condenser - for curbing evaporative loss in the vessel by cooling the exhaust gas path.</li>
		<li>Reagent pump lines for pH control or feed addition.</li>
	</ol>
	</li>
	<li>Clean the vessel and headplate with soap and water. A soft brush is recommended for scrubbing the vessel and headplate. Bleach or cleaners with chlorine cannot be used to clean the vessel.</li>
	<li>If the media is not heat labile, add it to the clean vessel. Only add until maximum working volume is reached. In this instance, maximum working volume is 3.3 L for a 5 L total volume vessel.</li>
	<li>Mount the vessel headplate, ensuring that the o-ring seal is properly seated.</li>
	<li>Install the exhaust gas condenser.</li>
	<li>Install the antifoam probe into its 10 mm port.</li>
	<li>Install the harvest pipe into its 10 mm port. Put a piece of tubing on the top of it and clamp it off.</li>
	<li>Install tubing and a 0.20 &#956;m filter on the sparge inlet and then clamp this off.</li>
	<li>Calibrate the pH probe:
	<ol>
		<li>Gather 2 reference buffers in small beakers, a wash bottle, and a spare wash beaker.</li>
		<li>Hook the pH probe up to the fermentor and turn the fermentor on.</li>
		<li>Scroll to the pH parameter and using the menu button, scroll to the &#39;Cal&#39; option, and press &#39;Enter&#39;.</li>
		<li>Select &#39;2&#39; for 2 point calibration.</li>
		<li>Wash off the probe tip and submerge in the low reference buffer. The value on the fermentor will stabilize. Using the + and - keys, adjust the displayed value to match the value of the pH reference buffer. Once it stops blinking, press &#39;Enter&#39;.</li>
		<li>Wash the probe off and insert it into the second reference buffer. Allow the value to stabilize and then use the keypad to make the displayed value match the high reference buffer value. Press &#39;Enter&#39; to confirm.</li>
		<li>Wash the probe tip off and disconnect it from the fermentor. Put the protective cap on the connection and install it in the headplate.</li>
	</ol>
	</li>
	<li>Open the pO2 probe tip and check to ensure that there is enough electrolyte to cover the tip. If not, add some to the reservoir and close it back up.</li>
	<li>Install the pO2 probe in the headplate and make sure that the autoclave cap is put on to protect its electrical connections. It will be calibrated after sterilization.</li>
	<li>Obtain a bottle for reagent feed with a dip tube and air filter. Add pump tubing to the dip tube port and attach the other end to the inlet port on the fermentor.</li>
	<li>Place tubing on all unused ports then clamp tubing off.</li>
	<li>Install a 0.45 &#956;m filter on the exhaust gas condenser. This filter ensures that the vessel does not pressurize in the autoclave.</li>
	<li>Check that all tubes that go below the level of the media are clamped off to prevent media from coming out.</li>
	<li>Put the vessel in the autoclave for 25 - 30 min at 121 &#176;C, liquid cycle. Caution: When it comes out it will be very hot.</li>
	<li>Install the vessel on the fermentor base.</li>
	<li>Hook up the pH probe cable.</li>
	<li>Attach the pO2 probe cable.</li>
	<li>Hook the sparger up to the gas addition rotameter.</li>
	<li>Sterilely add 1 M NaOH reagent to the bottle with the dip tube and install the pump head on the base pump head spindle.</li>
	<li>Put the temperature sensor in the thermowell on the headplate. Ensure that it goes all of the way to the bottom of the thermowell.</li>
	<li>Lower the agitation arm onto the vessels impeller coupling.</li>
	<li>Make sure that the fermentor is on.</li>
	<li>Check that water is available to the fermentor.</li>
	<li>Turn on the air supply to the fermentor.</li>
	<li>Scroll to the temperature parameter and set the temperature to 37 &#176;C. Press the &#39;Enter&#39; button to start the temperature control. The vessel will heat up in 15 min or less.</li>
	<li>Turn the gas flow to 1 vessel volume of media per minute or less. For this set up, gas flow is 3 L/min. Bubbles should appear at the bottom.</li>
	<li>Scroll to the agitation parameter and set to 300 rpm. Press &#39;Enter&#39; to turn on the agitation.</li>
	<li>Once the temperature has reached 37 &#176;C, scroll to the pH parameter and set it to 6.8. Turn the pH control on by pressing the &#39;Enter&#39; button.</li>
	<li>Set agitation to the maximum speed of 1,000 rpm for the run.</li>
	<li>Ensure that the pO2 probe is polarized properly to display proper values. After 2 hr, scroll to the pO2 parameter and use the &#39;Menu&#39; button to select the &#39;Calibrate Function&#39; and select the 1 point calibration.</li>
	<li>After 15 min, use the cursors to set the value to 100 and press &#39;Enter.&#39; This will calibrate pO2.</li>
	<li>Scroll to agitation and set to 300 rpm.</li>
	<li>Using the &#39;Menu&#39; button, turn the &#39;Cascade&#39; to &#39;on&#39; in the agitation menu. This will increase the speed of agitation in an effort to chop up air bubbles and force more oxygen into solution as growth demand increases.</li>
	<li>Scroll to the pO2 parameter and set the value to 30. Press the &#39;Enter&#39; button to start pO2.</li>
	<li>Start the control software package on the PC.</li>
	<li>Once the probes are calibrated, agitation is stable at 300 rpm, temperature is at 37 &#176;C and pH is close to 6.8, it is time to inoculate. Using alcohol, sterilize the port that will be used for inoculation.</li>
	<li>Draw the inoculum into a syringe and add it into the sterilized port. Close the port.</li>
	<li>Mark the time of inoculation in a log book.</li>
	<li>It is also important to take a sample at sterilization time. Sterilize the end of the harvest pipe tube with alcohol.</li>
	<li>Open the clamp covering the harvest port and use a syringe to draw a sample. This first sample is usually discarded because it has been sitting in the pipe.</li>
	<li>Use another syringe to pull another sample.</li>
	<li>With a third syringe, push air back through the pipe to remove as much dead volume from the line and clamp the line off.</li>
	<li>Determine the cell density and pH and record the values. With microbial cultures, it is useful to log values every hour. Interval is culture dependent.</li>
	<li>Harvest methodology depends upon the intent for the culture after growth is completed. In this case, sample the culture a final time to receive endpoint values for cell density and possible pH or measurement of wet cell weight. Open the headplate disposed of the contents in a designated &#34;kill tank&#34; with bleach or other antimicrobial agents.</li>
</ol>

<ol>
	<li>Shuler, M., Kargi, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Bioprocess Engineering Basic Concepts. , (2002).</li><li>Hambor, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioreactor+Design+and+Bioprocess+Controls+for+Industrialized+Cell+Processing.">Bioreactor Design and Bioprocess Controls for Industrialized Cell Processing.</a> BioProcess International. 10 (6), 22-33 (2012).</li><li>Julien, C., Whitford, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioreactor,+Monitoring,+Modeling+and+Simulation.">Bioreactor, Monitoring, Modeling and Simulation.</a> BioProcess International. 5 (01), S10-S17 (2007).</li><li>Fike, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nutrient+Supplementation+Strategies+for+Biopharmaceutical+Production.+Part+1:+Identifying+a+Formulation.">Nutrient Supplementation Strategies for Biopharmaceutical Production. Part 1: Identifying a Formulation.</a> BioProcess International. 7 (11), 46-52 (2009).</li><li>Jana, S., Deb, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategies+for+efficient+production+of+heterologous+proteins+in+Escherichia+coli.">Strategies for efficient production of heterologous proteins in Escherichia coli.</a> Appl Microbiol Biotechnol. 67, 289-298 (2005).</li><li>Wang, Z., Ly, M., Zhang, F., Zhong, W., Suen, A., Hickey, A. M., Dordick, J. S., Linhardt, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=E.+coli+K5+fermentation+and+the+preparation+of+heparosan,+a+bioengineered+heparin+precursor.">E. coli K5 fermentation and the preparation of heparosan, a bioengineered heparin precursor.</a> Biotechnol. Bioeng. 107, 964-973 (2010).</li><li>Baltz, R. H., Demain, A. L., Davies, J. 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> Manual of Industrial Microbiology and Biotechnology. , (2010).</li><li>Infors, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Minifors operating Manual and user guide. , .</li></ol>

Author A. Magno
 is an employee of ATR Biotech that produces reagents and instruments used in this article.

Stirred tank bioreactors are the standard in the Biotechnology industry and have been used for over 40 years 1. The small stirred tank has been important for scale-up, scale-down, strain optimization, characterization, and process development. It may also have an important role in the development of individualized medicine 2. The small scale bioreactor is most similar to in situ conditions for cell growth because it can be monitored and optimized throughout a run. Most often, initial experiments are performed using shake flasks but the conditions in the small scale bioreactor differ significantly from the shake flask. In one experiment we found that the conditions for optimal growth of E. coli and production of Green Fluorescent Protein (GFP) in the shake flask did not translate to the stirred tank (unpublished data).
Other methods of growing cells in large scale include roller bottles, single use rocking platform bioreactors 3 and larger single use bioreactors with working volumes from 50 to 5,000 L. Each method provides challenges for scale-up, but has found a place in production. The single use rocking platform bioreactor is similar to the stirred tank, and provides a regulated environment. It differs from the stirred tank in that mixing occurs due a rocking motion to generate waves to prevent cell settling and provide oxygenation. The hydrodynamics for this method are different from the stirred tank and maximum volume is limited to 1,000 L. The differences can affect cell growth and product production. Other single use systems combine the stirred tank with the disposable reactor to, provide a platform with a minimum of infrastructure and associated overhead, and the capability for high-throughput bioprocessing 4.
New users of benchtop Bioreactors may have trouble determining initial setpoints for pH, pO2 and temperature; however, published research can be referenced for this information 5, 6, 7, 8, 9. With bacterial cultures in particular, it is recommended to start agitation at the same speed as the shaker flask and the temperature at the same setpoint. Culture pH from previous shake flasks runs can also be used as a starting point. Setting of the pO2 value is more difficult and is typically determined empirically. However, beginning with 50% pO2 is a recommended starting point.

Basic fermentation technology is an extension of the simple shake flask technique for growing cultures. It grew out of the desire to control growth environments for live cultures in a more complete and quantitative way. Batch culture shake flasks are usually limited by imprecise control of temperature. Temperature uniformity in an incubated shaker or warm room is highly variable, sometimes straying 5 &#176;C or more from the intended setpoint. Since the shake flask is normally agitated at a fixed speed, oxygen uptake and gas exchange is limited. Once the available ambient oxygen is depleted, most cultures fail to thrive. There is no pH control in shake flasks. In many cases, if the culture is not limited by feed stock, it becomes acidic to the point of detriment to the culture and respiration slows dramatically. Most shake flask cultures are also run as a &#39;batch&#39;, which means that they are fed only once at or near the beginning of the cultures inoculation. After this initial carbon source is consumed, the culture stops growing. In some cases its metabolism may shift and begin to consume other metabolites in the culture broth, sometimes changing the characteristics of the resultant biomass or protein. Shake flasks are also usually subject to media evaporative loss in warmer culture environments, typically 10% of volume per 24 hr at 37 &#176;C. This loss changes the density of the culture and prohibits longer term operation of the system. Finally, the user may encounter foaming from the media after agitation. The occurrence of foam in the headspace above the culture will limit gas exchange and further stifle growth.
The basic fermentation system is designed to address all of these limitations. Careful temperature control is achieved in fermentation vessels by the use of impeller agitation and a heating jacket. A sensor inserted into the vessel and feedback control of heating and cooling of this jacket usually results in temperature control &#177;0.1 &#176;C around the setpoint. Benchtop fermentors generally provide control of pH via liquid reagent addition through a pump. The pH value is continuously monitored in an effort to keep the environment optimal for cell growth. Proper aeration is maintained by the aforementioned mixing impeller or by the infusion of air or oxygen supplemented gas directly into the culture. With shear-sensitive cultures, oxygen supplemented gas is the primary mechanism for maintenance of oxygen level in the culture. Measurement of oxygen in solution is usually achieved by a polarographic probe which is not normally available for use in shake flasks. It is also possible to continuously or periodically add feed to the vessel to maintain growth in a linear or exponential fashion. The exit gas condenser provides a cold surface for vapor in the exhaust gas flow to condense, thus preserving culture volume and density. Periodic addition of antifoam surfactant is actuated by a conductivity probe in the culture, reducing foam on the surface and allowing gas exchange.
The vessel, with all probes, fittings, impellers, harvest pipes, and tubes, is assembled and sterilized in a standard autoclave. After final probe calibrations and stabilization to operating environment, the culture is added to the vessel. The system can then be used to characterize the culture in a way that is more quantitative and precise than with a shake flask method. Tight control of temperature, pH, oxygen content, feed consumption, liquid evaporation, and foam levels all contribute to a much higher biomass and better protein yield.

<table border="0" cellpadding="0" cellspacing="0" width="356">
	<colgroup>
		<col />
		<col span="2" />
	</colgroup>
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;">Name</td>
			<td style="width:117px;">Company</td>
			<td style="width:117px;">Catalog Number</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;">LB broth</td>
			<td style="width:117px;">Sigma</td>
			<td style="width:117px;">L3022</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;">ampicillin</td>
			<td style="width:117px;">Sigma</td>
			<td style="width:117px;">A1593-25G</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;">LB broth</td>
			<td style="width:117px;">Sigma</td>
			<td style="width:117px;">L3022</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;">antifoam 204</td>
			<td style="width:117px;">Sigma</td>
			<td style="width:117px;">A8311</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;"> </td>
			<td style="width:117px;"> </td>
			<td style="width:117px;"> </td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">1 M Sodium Hydroxide</td>
			<td style="width:117px;">Sigma</td>
			<td>38215-1EA-R</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">Reference Buffer, pH 4.00</td>
			<td style="width:117px;">Sigma</td>
			<td style="width:117px;">B5020</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">Reference Buffer, pH 7.00</td>
			<td style="width:117px;">Sigma</td>
			<td style="width:117px;">B4770</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">pO2 probe electrolyte</td>
			<td style="width:117px;">Broadley James</td>
			<td>AS-3140-C30-0025</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;">0.45 micron filter</td>
			<td style="width:117px;">Cole Parmer</td>
			<td style="width:117px;">EW-02915-22</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:123px;">0.22 micron filter</td>
			<td style="width:117px;">Cole Parmer</td>
			<td style="width:117px;">EW-29950-40</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">Luer Lock syringe, 10 mL</td>
			<td style="width:117px;">Cole Parmer</td>
			<td style="width:117px;">EW-07940-12</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">Minifors Bioreactor</td>
			<td style="width:117px;">Infors HT</td>
			<td style="width:117px;">B-Pack 5.0</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">Air Admiral air pump</td>
			<td style="width:117px;">Cole Parmer</td>
			<td style="width:117px;">EW-79202-00</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:123px;">1175 PD Chilling circulator</td>
			<td style="width:117px;">VWR</td>
			<td style="width:117px;">13721-204</td>
		</tr>
	</tbody>
</table>

operation of a benchtop bioreactor

Fermentation systems are used to provide an optimal growth environment for many different types of cell cultures. The ability afforded by fermentors to carefully control temperature, pH, and dissolved oxygen concentrations in particular makes them essential to efficient large scale growth and expression of fermentation products. This video will briefly describe the advantages of the fermentor over the shake flask. It will also identify key components of a typical benchtop fermentation system and give basic instruction on setup of the vessel and calibration of its probes. The viewer will be familiarized with the sterilization process and shown how to inoculate the growth medium in the vessel with culture. Basic concepts of operation, sampling, and harvesting will also be demonstrated. Simple data analysis and system cleanup will also be discussed.

The bioreactor can be run with or without the IRIS software. However to capture data, it is best to use the software. Before addition of bacteria, the pH and oxygen sensors must be calibrated, the impeller speed set and the temperature set. In Figure 2, the data output for a bioreactor run is presented. The temperature was set to 30 &#176;C, the impeller speed at 200 rpm. The parameters may be different for each experiment but before addition of bacteria, the system should be at steady state. In Figure 3, the change in oxygen concentration with the addition of the bacterial culture is shown. The set-points for this experiment are 37 &#176;C, stirrer at 200 rpm, O2 at 70%. At time 19:20, the feed pump delivers the bacterial seed culture at an OD of 0.1 causing an immediately drop in the O2 level. The bioreactor responds to the changing O2 level with an increase in the airflow and impeller speed. The set-points for the increases are set in the cascade (step 35). The reactor pH was monitored in real time, with the pH decreasing then increasing as demonstrated by the pH line fluctuation over time (Figure 4). The entire run can be analyzed and parameters adjusted for subsequent experiments.
<img alt="Figure 1" src="/files/ftp_upload/50582/50582fig1.jpg" /> 
Figure 1. Benchtop bioreactor with all parts labeled and connected. This figure shows the stirred tank reactor at the beginning of a run. The parts of the reactor are labeled and include the sensors, impeller, temperature gauge, feed &#38; sampling bottles, exhaust port, air pressure gauge, heating jacket, condenser tower, and control panel.
<img alt="Figure 2" src="/files/ftp_upload/50582/50582fig2.jpg" /> 
Figure 2. Screen image of bioreactor software at the beginning of run. At the start of a stirred tank run, the temperature was set at 30 &#176;C (yellow line), impeller speed at 200 rpm (red line), pH at 6.5 (green line), and pO2 at 57.2% (blue line). <a href="https://www.jove.com/files/ftp_upload/50582/50582fig2large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/50582/50582fig3.jpg" /> 
Figure 3. Screen image of bioreactor software during the run as oxygen decreases. When the culture is in active growth phase, it consumes oxygen and the amount of oxygen in the reactor is decreased (blue line). By increasing the impeller speed, more oxygen can be added to the culture (red line). <a href="https://www.jove.com/files/ftp_upload/50582/50582fig3large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 4" src="/files/ftp_upload/50582/50582fig4.jpg" /> 
Figure 4. Screen image of bioreactor software showing pH change over time. The pH of the culture is monitored continuously and over time it decreases as lactic acid is produced and then increases as base is added to the bioreactor (blue line). <a href="https://www.jove.com/files/ftp_upload/50582/50582fig4large.jpg" target="_blank"> Click here to view larger figure</a>.

Watch this Scientific Journal Video about Operation of a Benchtop Bioreactor at JoVE.com

Operation of a Benchtop Bioreactor

Fermentors are used to increase culture yield and productivity of bioengineered cells. After screening multiple microbial or animal cell culture candidates in shake flasks, the next logical step is to increase the selected culture&#x2019;s biomass with the fermentor. This video demonstrates the setup and operation of a typical benchtop bioreactor system.

Fermentation systems are used to provide an optimal growth environment for many different types of cell cultures. The ability afforded by fermentors to ...

Research

JoVE Journal

Bioengineering

Engineering Biological-Based Vascular Grafts Using a Pulsatile Bioreactor

Much effort has been devoted to develop and advance the methodology to regenerate functional small-diameter arterial bypasses. In the physiological environment, both mechanical and chemical stimulation are required to maintain the proper development and functionality of arterial vessels1,2.
Bioreactor culture systems developed by our group are designed to support vessel regeneration within a precisely controlled chemo-mechanical environment mimicking that of native vessels. Our bioreactor assembly and maintenance procedures are fairly simple and highly repeatable3,4. Smooth muscle cells (SMCs) are seeded onto a tubular polyglycolic acid (PGA) mesh that is threaded over compliant silicone tubing and cultured in the bioreactor with or without pulsatile stimulation for up to 12 weeks. There are four main attributes that distinguish our bioreactor from some predecessors. 1) Unlike other culture systems that simulate only the biochemical surrounding of native blood vessels, our bioreactor also creates a physiological pulsatile environment by applying cyclic radial strain to the vessels in culture. 2) Multiple engineered vessels can be cultured simultaneously under different mechanical conditions within a controlled chemical environment. 3) The bioreactor allows a mono layer of endothelial cells (EC) to be easily coated onto the luminal side of engineered vessels for animal implantation models. 4) Our bioreactor can also culture engineered vessels with different diameter size ranged from 1 mm to 3 mm, saving the effort to tailor each individual bioreactor to fit a specific diameter size. 
The engineered vessels cultured in our bioreactor resemble native blood vessels histologically to some degree. Cells in the vessel walls express mature SMC contractile markers such as smooth muscle myosin heavy chain (SMMHC)3. A substantial amount of collagen is deposited within the extracellular matrix, which is responsible for ultimate mechanical strength of the engineered vessels5. Biochemical analysis also indicates that collagen content of engineered vessels is comparable to that of native arteries6. Importantly, the pulsatile bioreactor has consistently regenerated vessels that exhibit mechanical properties that permit successful implantation experiments in animal models3,7. Additionally, this bioreactor can be further modified to allow real-time assessment and tracking of collagen remodeling over time, non-invasively, using a non-linear optical microscopy (NLOM)8. To conclude, this bioreactor should serve as an excellent platform to study the fundamental mechanisms that regulate the regeneration of functional small-diameter vascular grafts.

Our group has developed a bioreactor culture system that mimics the physiological pulsatile stresses of the cardiovascular system to regenerate implantable small-diameter vascular grafts.

Much effort has been devoted to develop and advance the methodology to regenerate functional small-diameter arterial bypasses. In the physiological ...

Design of a Biaxial Mechanical Loading Bioreactor for Tissue Engineering

We designed a loading device that is capable of applying uniaxial or biaxial mechanical strain to a tissue engineered biocomposites fabricated for transplantation. While the device primarily functions as a bioreactor that mimics the native mechanical strains, it is also outfitted with a load cell for providing force feedback or mechanical testing of the constructs. The device subjects engineered cartilage constructs to biaxial mechanical loading with great precision of loading dose (amplitude and frequency) and is compact enough to fit inside a standard tissue culture incubator. It loads samples directly in a tissue culture plate, and multiple plate sizes are compatible with the system. The device has been designed using components manufactured for precision-guided laser applications. Bi-axial loading is accomplished by two orthogonal stages. The stages have a 50 mm travel range and are driven independently by stepper motor actuators, controlled by a closed-loop stepper motor driver that features micro-stepping capabilities, enabling step sizes of less than 50 nm. A polysulfone loading platen is coupled to the bi-axial moving platform. Movements of the stages are controlled by Thor-labs Advanced Positioning Technology (APT) software. The stepper motor driver is used with the software to adjust load parameters of frequency and amplitude of both shear and compression independently and simultaneously. Positional feedback is provided by linear optical encoders that have a bidirectional repeatability of 0.1 &mu;m and a resolution of 20 nm, translating to a positional accuracy of less than 3 &mu;m over the full 50 mm of travel. These encoders provide the necessary position feedback to the drive electronics to ensure true nanopositioning capabilities. In order to provide the force feedback to detect contact and evaluate loading responses, a precision miniature load cell is positioned between the loading platen and the moving platform. The load cell has high accuracies of 0.15% to 0.25% full scale.

We designed a novel mechanical loading bioreactor that can apply uniaxial or biaxial mechanical strain to a cartilage biocomposite prior to transplantation into an articular cartilage defect.

We  designed a loading device that is capable of applying uniaxial or biaxial  mechanical strain to a tissue engineered biocomposites fabricated for  ...

Microfluidic Picoliter Bioreactor for Microbial Single-cell Analysis: Fabrication, System Setup, and Operation

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described in detail. The PLBR can be utilized for the analysis of single bacteria and microcolonies to investigate biotechnological and microbiological related questions concerning, e.g.&#160;cell growth, morphology, stress response, and metabolite or protein production on single-cell level. The device features continuous media flow enabling constant environmental conditions for perturbation studies, but in addition allows fast medium changes as well as oscillating conditions to mimic any desired environmental situation. To fabricate the single use devices, a silicon wafer containing sub micrometer sized SU-8 structures served as the replication mold for rapid polydimethylsiloxane casting. Chips were cut, assembled, connected, and set up onto a high resolution and fully automated microscope suited for time-lapse imaging, a powerful tool for spatio-temporal cell analysis. Here, the biotechnological platform organism Corynebacterium glutamicum&#160;was seeded into the PLBR and cell growth and intracellular fluorescence were followed over several hours unraveling time dependent population heterogeneity on single-cell level, not possible with conventional analysis methods such as flow cytometry. Besides insights into device fabrication, furthermore, the preparation of the preculture, loading, trapping of bacteria, and the PLBR cultivation of single cells and colonies is demonstrated. These devices will add a new dimension in microbiological research to analyze time dependent phenomena of single bacteria under tight environmental control. Due to the simple and relatively short fabrication process the technology can be easily adapted at any microfluidics lab and simply tailored towards specific needs.

In this protocol the fabrication, setup and basic operation of a microfluidic picoliter bioreactor (PLBR) for single-cell analysis of prokaryotic microorganisms is introduced. Industrially relevant microorganisms were analyzed as proof of principle allowing insights into growth rate, morphology, and phenotypic heterogeneity over certain time periods, hardly possible with conventional methods.

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described ...

Construction and Characterization of a Novel Vocal Fold Bioreactor

In vitro engineering of mechanically active tissues requires the presentation of physiologically relevant mechanical conditions to cultured cells. To emulate the dynamic environment of vocal folds, a novel vocal fold bioreactor capable of producing vibratory stimulations at fundamental phonation frequencies is constructed and characterized. The device is composed of a function generator, a power amplifier, a speaker selector and parallel vibration chambers. Individual vibration chambers are created by sandwiching a custom-made silicone membrane between a pair of acrylic blocks. The silicone membrane not only serves as the bottom of the chamber but also provides a mechanism for securing the cell-laden scaffold. Vibration signals, generated by a speaker mounted underneath the bottom acrylic block, are transmitted to the membrane aerodynamically by the oscillating air. Eight identical vibration modules, fixed on two stationary metal bars, are housed in an anti-humidity chamber for long-term operation in a cell culture incubator. The vibration characteristics of the vocal fold bioreactor are analyzed non-destructively using a Laser Doppler Vibrometer (LDV). The utility of the dynamic culture device is demonstrated by culturing cellular constructs in the presence of 200-Hz sinusoidal vibrations with a mid-membrane displacement of 40 &#181;m. Mesenchymal stem cells cultured in the bioreactor respond to the vibratory signals by altering the synthesis and degradation of vocal fold-relevant, extracellular matrix components. The novel bioreactor system presented herein offers an excellent in vitro platform for studying vibration-induced mechanotransduction and for the engineering of functional vocal fold tissues.

A novel vocal fold bioreactor capable of delivering physiologically relevant, vibratory stimulation to cultured cells is constructed and characterized. This dynamic culture device, when combined with a fibrous poly(&#949;-caprolactone) scaffold, creates a vocal fold-mimetic environment that modulates the behaviors of mesenchymal stem cells.

In vitro engineering of mechanically active tissues requires the presentation of physiologically relevant mechanical conditions to cultured cells. To ...

Cultivation of Mammalian Cells Using a Single-use Pneumatic Bioreactor System

Recent advances in mammalian, insect, and stem cell cultivation and scale-up have created tremendous opportunities for new therapeutics and personalized medicine innovations. However, translating these advances into therapeutic applications will require in vitro systems that allow for robust, flexible, and cost effective bioreactor systems. There are several bioreactor systems currently utilized in research and commercial settings; however, many of these systems are not optimal for establishing, expanding, and monitoring the growth of different cell types. The culture parameters most challenging to control in these systems include, minimizing hydrodynamic shear, preventing nutrient gradient formation, establishing uniform culture medium aeration, preventing microbial contamination, and monitoring and adjusting culture conditions in real-time. Using a pneumatic single-use bioreactor system, we demonstrate the assembly and operation of this novel bioreactor for mammalian cells grown on micro-carriers. This bioreactor system eliminates many of the challenges associated with currently available systems by minimizing hydrodynamic shear and nutrient gradient formation, and allowing for uniform culture medium aeration. Moreover, the bioreactor&#8217;s software allows for remote real-time monitoring and adjusting of the bioreactor run parameters. This bioreactor system also has tremendous potential for scale-up of adherent and suspension mammalian cells for production of a variety therapeutic proteins, monoclonal antibodies, stem cells, biosimilars, and vaccines.

Using a pneumatic bioreactor, we demonstrate the assembly, operation, and performance of this single-use bioreactor system for the growth of mammalian cells.

Recent advances in mammalian, insect, and stem cell cultivation and scale-up have created tremendous opportunities for new therapeutics and personalized ...

A Novel Bioreactor for High Density Cultivation of Diverse Microbial Communities

A novel reactor design, coined a high density bioreactor (HDBR), is presented for the cultivation and study of high density microbial communities. Past studies have evaluated the performance of the reactor for the removal of COD1 and nitrogen species2-4 by heterotrophic and chemoautotrophic bacteria, respectively. The HDBR design eliminates the requirement for external flocculation/sedimentation processes while still yielding effluent containing low suspended solids. In this study, the HDBR is applied as a photobioreactor (PBR) in order to characterize the nitrogen removal characteristics of an algae-based photosynthetic microbial community. As previously reported for this HDBR design, a stable biomass zone was established with a clear delineation between the biologically active portion of the reactor and the recycling reactor fluid, which resulted in a low suspended solid effluent. The algal community in the HDBR was observed to remove 18.4% of total nitrogen species in the influent. Varying NH4+ and NO3- concentrations in the feed did not have an effect on NH4+ removal (n=44, p=0.993 and n=44, p=0.610 respectively) while NH4+ feed concentration was found to be negatively related with NO3- removal (n=44, p=0.000) and NO3- feed concentration was found to be positively correlated with NO3- removal (n=44, p=0.000). Consistent removal of NH4+, combined with the accumulation of oxidized nitrogen species at high NH4+ fluxes indicates the presence of ammonia- and nitrite-oxidizing bacteria within the microbial community.

A novel reactor design, coined a high density bioreactor (HDBR), is presented for the cultivation and study of high density microbial communities. Here, the HDBR is successfully applied in a photobioreactor (PBR) configuration for the study of nitrogen metabolism by a mixed high density algal community.

A novel reactor design, coined a high density bioreactor (HDBR), is presented for the cultivation and study of high density microbial communities. Past ...

Assembly and Operation of an Acoustofluidic Device for Enhanced Delivery of Molecular Compounds to Cells

Efficient intracellular delivery of biomolecules is required for a broad range of biomedical research and cell-based therapeutic applications. Ultrasound-mediated sonoporation is an emerging technique for rapid intracellular delivery of biomolecules. Sonoporation occurs when cavitation of gas-filled microbubbles forms transient pores in nearby cell membranes, which enables rapid uptake of biomolecules from the surrounding fluid. Current techniques for in vitro sonoporation of cells in suspension are limited by slow throughput, variability in the ultrasound exposure conditions for each cell, and high cost. To address these limitations, a low-cost acoustofluidic device has been developed which integrates an ultrasound transducer in a PDMS-based fluidic device to induce consistent sonoporation of cells as they flow through the channels in combination with ultrasound contrast agents. The device is fabricated using standard photolithography techniques to produce the PDMS-based fluidic chip. An ultrasound piezo disk transducer is attached to the device and driven by a microcontroller. The assembly can be integrated inside a 3D-printed case for added protection. Cells and microbubbles are pushed through the device using a syringe pump or a peristaltic pump connected to PVC tubing. Enhanced delivery of biomolecules to human T cells and lung cancer cells is demonstrated with this acoustofluidic system. Compared to bulk treatment approaches, this acoustofluidic system increases throughput and reduces variability, which can improve cell processing methods for biomedical research applications and manufacturing of cell-based therapeutics.

This protocol describes the assembly and operation of a low-cost acoustofluidic device for rapid molecular delivery to cells via sonoporation induced by ultrasound contrast agents.

Efficient intracellular delivery of biomolecules is required for a broad range of biomedical research and cell-based therapeutic applications. ...

Open-source Toolkit: Benchtop Carbon Fiber Microelectrode Array for Nerve Recording

Conventional peripheral nerve probes are primarily fabricated in a cleanroom, requiring the use of multiple expensive and highly specialized tools. This paper presents a cleanroom "light" fabrication process of carbon fiber neural electrode arrays that can be learned quickly by an inexperienced cleanroom user. This carbon fiber electrode array fabrication process requires just one cleanroom tool, a Parylene C deposition machine, that can be learned quickly or outsourced to a commercial processing facility at marginal cost. This fabrication process also includes hand-populating printed circuit boards, insulation, and tip optimization. 
The three different tip optimizations explored here (Nd:YAG laser, blowtorch, and UV laser) result in a range of tip geometries and 1 kHz impedances, with blowtorched fibers resulting in the lowest impedance. While previous experiments have proven laser and blowtorch electrode efficacy, this paper also shows that UV laser-cut fibers can record neural signals in vivo. Existing carbon fiber arrays either do not have individuated electrodes in favor of bundles or require cleanroom fabricated guides for population and insulation. The proposed arrays use only tools that can be used at a benchtop for fiber population. This carbon fiber electrode array fabrication process allows for quick customization of bulk array fabrication at a reduced price compared to commercially available probes.

Here, we describe fabrication methodology for customizable carbon fiber electrode arrays for recording in vivo in nerve and brain.

Conventional peripheral nerve probes are primarily fabricated in a cleanroom, requiring the use of multiple expensive and highly specialized tools. This ...

The Fabrication and Operation of a Continuous Flow, Micro-Electroporation System with Permeabilization Detection

Current therapeutic innovations, such as CAR-T cell therapy, are heavily reliant on viral-mediated gene delivery. Although efficient, this technique is accompanied by high manufacturing costs, which has brought about an interest in using alternative methods for gene delivery. Electroporation is an electro-physical, non-viral approach for the intracellular delivery of genes and other exogenous materials. Upon the application of an electric field, the cell membrane temporarily allows molecular delivery into the cell. Typically, electroporation is performed on the macroscale to process large numbers of cells. However, this approach requires extensive empirical protocol development, which is costly when working with primary and difficult-to-transfect cell types. Lengthy protocol development, coupled with the requirement of large voltages to achieve sufficient electric-field strengths to permeabilize the cells, has led to the development of micro-scale electroporation devices. These micro-electroporation devices are manufactured using common microfabrication techniques and allow for greater experimental control with the potential to maintain high throughput capabilities. This work builds off a microfluidic-electroporation technology capable of detecting the level of cell membrane permeabilization at a single-cell level under continuous flow. However, this technology was limited to 4 cells processed per second, and thus a new approach for increasing the system throughput is proposed and presented here. This new technique, denoted as cell-population-based feedback control, considers the cell permeabilization response to a variety of electroporation pulsing conditions and determines the best-suited electroporation pulse conditions for the cell type under test. A higher-throughput mode is then used, where this 'optimal' pulse is applied to the cell suspension in transit. The steps for fabricating the device, setting up and running the microfluidic experiments, and analyzing the results are presented in detail. Finally, this micro-electroporation technology is demonstrated by delivering a DNA plasmid encoding for green fluorescent protein (GFP) into HEK293 cells.

This protocol describes the microfabrication techniques required to build a lab-on-a-chip, microfluidic electroporation device. The experimental setup performs controlled, single-cell-level transfections in a continuous flow and can be extended to higher throughputs with population-based control. An analysis is provided showcasing the ability to electrically monitor the degree of cell membrane permeabilization in real-time.

Current therapeutic innovations, such as CAR-T cell therapy, are heavily reliant on viral-mediated gene delivery. Although efficient, this technique is ...

Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue

Repeated injury to airway tissue can impair lung function and cause chronic lung disease, such as chronic obstructive pulmonary disease. Advances in regenerative medicine and bioreactor technologies offer opportunities to produce lab-grown functional tissue and organ constructs that can be used to screen drugs, model disease, and engineer tissue replacements. Here, a miniaturized bioreactor coupled with an imaging modality that allows in situ visualization of the inner lumen of explanted rat trachea during in vitro tissue manipulation and&#160;culture is described. Using this bioreactor, the protocol demonstrates imaging-guided selective removal of endogenous cellular components while preserving the intrinsic biochemical features and ultrastructure of the airway tissue matrix. Furthermore, the delivery, uniform distribution, and subsequent prolonged culture of exogenous cells on the decellularized airway lumen with optical monitoring in situ are shown. The results highlight that the imaging-guided bioreactor can potentially be used to facilitate the generation of functional in vitro airway tissues.

The protocol describes an imaging-enabled bioreactor that allows the selective removal of the endogenous epithelium from the rat trachea and homogenous distribution of exogenous cells on the lumen surface, followed by long-term in vitro culture of the cell-tissue construct.

Repeated injury to airway tissue can impair lung function and cause chronic lung disease, such as chronic obstructive pulmonary disease. Advances in ...