This study was supported by the Natural Sciences and Engineering Research Council (NSERC), Canada Foundation for Innovation (CFI), Canada First Research Excellence Fund (CFREF), Alberta Innovates, General Sir John Monash Foundation, the Government of Alberta, and the University of Calgary. Thanks is extended to Mark Toonen for the electrical work and William Richardson for solubility calculations.

1. Construction of the PBR stand
<ol>
	<li>With a handheld hacksaw, cut five 380 mm lengths and two 200 mm lengths of angled slotted steel. Fasten together with bolts and large corner braces to make a stand&#39;s base (Figure 1A). Glue on safety end caps.</li>
	<li>Connect two uncut (1220 mm) vertical lengths of angled slotted steel to the base. Secure with bolts and metal corner gussets (Figure 1B). Glue on safety end caps.</li>
	<li>Cut four 65 mm flat lengths of slotted steel. Bolt these at 90&#176; angles onto the vertical supports&#8212;attach two to each support, one 130 mm up from the base (Figure 1C) and one 60 mm down from the top.</li>
	<li>Fasten vertical supports across their top with a horizontal 140 mm length of flat slotted steel (crossbeam) bolted to the back of the frame (Figure 2A).</li>
</ol>
2. Construction of the light chamber
<ol>
	<li>Cut a white 153 mm diameter polyvinyl chloride (PVC) pipe to a length of 1070 mm. Cut the pipe in half lengthwise with a bandsaw. Sand all edges.</li>
	<li>Evenly space and center four aluminum heat sink channels vertically along with the interior of the pipe. Do not attach channels within 20 mm from the pipe&#39;s cut edge. With small bolts, secure the channels in place at their top and bottom (Figure 2B).</li>
	<li>Bolt one-half of the pipe to the stand using the horizontal supports fashioned in step 1.3.</li>
	<li>Lay the reactor down and reunite the pipe halves by taping them together. Center the piano hinge along one cutline. Trace the hinge holes and drill the pipe accordingly. Use a rivet gun and medium-length rivets to fasten the hinge to the pipe.</li>
	<li>Use a small bungee cord (see Table of Materials) to hold the pipe shut (Figure 1C).</li>
	<li>Consult an electrician to wire in the LED lights and install the following four components: LED driver, digital multiplex (DMX) decoder, DMX lighting controller, and switch box (see Table of Materials). Fix all the components to the rear of the PBR according to Figure 2A.</li>
</ol>
3. Construction of the bottle platforms
<ol>
	<li>Cut platform shapes (Figure 3A) out of hard plastic, e.g., high density polyethylene (HDPE) (see Table of Materials), using computer numerical control (CNC) milling. Make three of each shape. 
	NOTE: Cutting spare top and bottom layers is recommended.</li>
	<li>Tape the bottom and top layer together. Mark and drill five small holes, 6 mm in diameter, through both shapes (Figure 3A). Using a larger drill bit, carefully expand the surface of these holes so that bolt heads can be recessed (Figure 3B).</li>
	<li>For each of the three platforms, bolt two small corner braces to the back half of the pipe. 
	NOTE: The distance between the top of the braces should be 350 mm.</li>
	<li>Center each bottom layer on top of its braces. Mark the location of the drill holes from beneath the braces. Drill two 6 mm diameter holes. Using a larger drill bit, carefully expand the surface of the holes so that bolts can be recessed.</li>
	<li>Bolt bottom layers to their braces (Figure 3B&#8211;C).</li>
	<li>Cut fifteen pieces of 12 mm long 6.35 mm outside diameter (OD) rigid tubing. Sandwich five pieces of the rigid tube between each top and bottom layer. Fasten layers and tubing together with long narrow bolts according to Figure 3B&#8211;D.</li>
	<li>Drill a large hole in the PVC pipe behind each platform. Insert each micro magnetic stirrer into its platform. Thread each stirrer&#39;s electrical cable through these newly cut holes (Figure 3C&#8211;E). Connect each stirrer to its respective control unit as well as a power outlet.</li>
	<li>Place a 1 L bottle on each platform. Add eye bolts in the rear pipe at bottle neck level. Wrap a small bungee cord around each bottle neck to add stability (Figure 4A). 
	&#8203;NOTE: Throughout the protocol, color coding platforms, bottles, magnetic stirrers, and all associated cables and sensors will be helpful.</li>
</ol>
4. Construction of the liquid sampling ports (optional)
<ol>
	<li>Cut three 60 mm lengths of 6.35 mm OD rigid tubing. Using a 5 mm drill bit, drill holes through each rubber stopper. Push rigid tube lengths through the stopper.</li>
	<li>Cut three 60 mm lengths of 3.18 mm OD rigid tubing. Connect these with a straight reducer to the protruding tube on the underside of each stopper.</li>
	<li>Insert a one-way stopcock valve (e.g., port 1 = female Luer, port 2 = male slip Luer) in the protruding tube on each stopper surface (Figure 4A).</li>
	<li>Cut three 30 mm lengths of 3.18 mm OD flexible tubing. Insert Luer fittings (e.g., male Luer to hose barb and female Luer to hose barb) on either end.</li>
	<li>Connect the pieces made in step 4.4 to the stopcock valves on the surface of each rubber stopper (Figure 4A). 
	&#8203;NOTE: Many combinations of stopcocks and Luer fittings can produce the same result. The design should allow liquid to be drawn out or inserted via a syringe.</li>
</ol>
5. Hooking up volumetric gas sensors
<ol>
	<li>Prepare gas sensors according to the manufacturer&#39;s instructions. 
	NOTE: This mainly involves filling gas sensors with packing liquid (Figure 4B).</li>
	<li>To make the gas lines, cut three 1000 mm lengths of 3.18 mm OD flexible tubing.</li>
	<li>Drill three 4 mm diameter holes in the rear PVC pipe. Position holes next to the hinge at bottle neck height. Thread gas lines through these holes (Figure 4A).</li>
	<li>To the end of the gas line inside the PVC pipe, add a Luer fitting (e.g., hose barb to male Luer) and connect a one-way stopcock valve (e.g., port 1 = female Luer, port 2 = male Luer). 
	NOTE: The valve is only required when liquid sampling ports are also installed.</li>
	<li>Join the other end of the gas line to the inlet port of the gas sensor using a straight reducer. Secure this connection with a zip tie.</li>
	<li>Connect all gas sensors to the digital input module (DIM) with jack plug cables and the DIM to a nearby computer.</li>
	<li>Install the data acquisition software (see Table of Materials) on a Windows operating system and plug in the licensing key dongle. Add sensor calibration files to the software&#39;s calibration directory.</li>
</ol>
6. Programming of the light regime
<ol>
	<li>Using the rear on/off switch, turn on the PBR and connect the DMX lighting controller to a computer via a micro-USB cable.</li>
	<li>Download Store Upgrade Tools (SUT) and the LED control software (see Table of Materials). Register the DMX lighting controller online.</li>
	<li>Open the LED control software and select Click here to work with the USB-DMX interface: SUSHI-RB-RJ.</li>
	<li>Under the Set-up tab in the ScanLibrary box, select the Generic folder and Single Channel. Change ScanLibrary settings to DMX universe 1, the number of fixtures to 4, and the index number to 1. In the upper-right corner, change the drop-down box to List View. Lastly, click on Patch (Supplementary Figure 1). 
	NOTE: In the DMX universe, one box tests each LED strip&#39;s control by sliding the dimmer buttons or entering a numerical value into the text box.</li>
	<li>Make a standard curve that relates the digital light setting in the lighting control software to the light intensity experienced in the center of the PVC tube (Figure 5). Measure internal light intensity with a small spherical probe (see Table of Materials) suspended in the center of the PVC pipe.</li>
	<li>Advance to the Editor tab. To build a custom light program, create a new Scene and begin adding Steps. Refer to Supplementary Table 1 for an example 16:8 h diurnal program. Set the scene to loop. 
	NOTE: Steps break down scenes into blocks of time, each of which can be set to different light intensity. Steps range from 1 s to 43 min. Here, 30 min steps are most convenient. Multiple scenes can be loaded onto one DMX lighting controller device.</li>
	<li>Create an additional helper scene that is immediately recognizable, e.g., two of the four LEDs on. 
	NOTE: Scenes can be cycled through manually using the button on the side of the DMX lighting controller. If the desired light program begins during the night, it will be impossible to distinguish whether the light program has already started. The helper scene serves as an indicator that the DMX lighting controller is functioning correctly.</li>
	<li>Save the scenes and advance to the Stand Alone tab. Write the memory of the DMX lighting controller and disconnect the device from the computer.</li>
	<li>Connect the DMX lighting controller to its power source using a micro-USB.</li>
	<li>Before starting an experiment, test the light program by logging the internal light intensity for a 24 h duration. If the liquid temperature is of interest, log this simultaneously with a submerged temperature probe (Figure 6).</li>
</ol>
7. Starting an experiment
<ol>
	<li>Sterilize media, bottles, stir bars, rubber stoppers, sampling ports, threaded aperture screw caps, and tubing. 
	NOTE: All components used in this design are autoclavable except the valves and Luer fittings&#8212;there are autoclavable alternatives from other manufacturers.</li>
	<li>Open the data acquisition software and fill in the configuration page (Supplementary Figure 2). Assign calibration files to their respective sensors.</li>
	<li>Under Directory File Name select the corresponding DIM port number folder. Click on Current Folder&#160;and repeat for all ports.</li>
	<li>Click on OK to move on to the logging page.</li>
	<li>Fill bottles to the desired volume with cultivation medium (Supplementary Tables 2,3). 
	NOTE: Each of these bottles will hold a maximum of ~1.1 L with a small headspace (~80 mL, including the gas line).</li>
	<li>Centrifuge the stock culture in three balanced 50 mL tubes for 15 min at 4500 x g to yield three pellets. Add one pellet to each bottle&#8211;wash in pellets with a serological pipette and fresh medium. 
	NOTE: The culture density of day 0 is also known as the initial biomass concentration (IBC). To measure the IBC in gAFDW.L-1, an additional 50 mL tube may be centrifuged in step 7.6. The resulting pellet can then be dried and combusted15,19. Step 7.6 will likely require modification based on the individual aims of users and their experiments.</li>
	<li>Drop a magnetic stirrer, 25 x 8 mm, into each bottle.</li>
	<li>Seal each bottle opening with a rubber stopper and threaded aperture screw cap (Figure 4A). If optional sampling ports are installed, close the valves.</li>
	<li>Locate the end of each gas line inside the PVC pipe (built in step 5.4) and attach a needle to the valve&#39;s male Luer port.</li>
	<li>Connect each bottle to its gas sensor by piercing each rubber stopper with the corresponding needle.</li>
	<li>Launch each gas sensor individually by checking the tick box&#160;on the left side of the screen, clicking on Start, and inputting a file name. Click OK and repeat for all sensors&#160;(Supplementary Figure 3). 
	NOTE: While logging, do not exit the data acquisition window. Set power and sleep settings of the computer to never and postpone computer updates for the duration of the experiment.</li>
	<li>Switch on the PBR and ensure that the DMX lighting controller is plugged into a power supply. The first programmed scene will automatically commence. Refer to step&#160;6.7&#160;to confirm that the DMX lighting controller is functioning correctly.</li>
</ol>
8. Bottle sampling (optional)
<ol>
	<li>Prepare an additional 500 mL of the fresh medium before commencing the experiment (Supplementary Table 2). 
	NOTE: If a 24 h light program with a 16:8 h diurnal cycle was started at 9 am, then sampling times before dusk and dawn would fall at 8 am and 4 pm (Supplementary Table 1). Here, dawn and dusk refer to 30 min steps that transition lights from ON to OFF and vice versa.</li>
	<li>Close the valve on the gas line.</li>
	<li>Connect a syringe (10 mL) to the sampling port valve (Figure 4A).</li>
	<li>Open the sampling port valve and withdraw 8 mL of culture. 
	NOTE: Between 5&#8211;10 mL is recommended. Removing liquid generates a vacuum in the headspace, making volumes &#62;10 mL difficult to extract.</li>
	<li>Close the sampling port valve and disconnect the syringe.</li>
	<li>Connect a syringe containing 8 mL of fresh medium (from step 8.1) to the sampling port valve.</li>
	<li>Open the sampling port valve and inject fresh medium. 
	NOTE: Replacing the volume of sampled culture with fresh medium serves to maintain an equal headspace volume and pressure and flush the sampling port line.</li>
	<li>Close the sampling port valve before disconnecting the syringe.</li>
	<li>Repeat steps 8.2&#8211;8.8 at each sampling time.</li>
</ol>
9. Ending an experiment
<ol>
	<li>Check all active port tick boxes in the data acquisition window and click on Stop.</li>
	<li>To export data, select File and Offline Data. Select all relevant log files. Export the data to spreadsheet software and save.</li>
	<li>For each bottle, convert the total volume of oxygen measured in mL to moles using the ideal gas law. Predict the weight of biomass grown (gAFDW) if 1.05 mole of O2 is generated for each mole of biomass produced. Take the molar weight of biomass as 24.6 g mol-1.</li>
	<li>Manually curate the flow rate data. Use units of mL/h and a 3-point moving average.</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+of+the+microalgae+Neochloris+oleoabundans+at+high+partial+oxygen+pressures+and+sub-saturating+light+intensity.">Growth of the microalgae Neochloris oleoabundans at high partial oxygen pressures and sub-saturating light intensity.</a> Bioresource Technology. 104, 565-570 (2012).</li><li>Latifi, A., Ruiz, M., Zhang, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxidative+stress+in+cyanobacteria.">Oxidative stress in cyanobacteria.</a> FEMS microbiology reviews. 33 (2), 258-278 (2009).</li><li>Morillas-Espa&#241;a, A., Lafarga, T., G&#243;mez-Serrano, C., Aci&#233;n-Fern&#225;ndez, F. G., Gonz&#225;lez-L&#243;pez, C. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Year-long+production+of+Scenedesmus+almeriensis+in+pilot-scale+raceway+and+thin-layer+cascade+photobioreactors.">Year-long production of Scenedesmus almeriensis in pilot-scale raceway and thin-layer cascade photobioreactors.</a> Algal Research. 51, (2020).</li><li>Melis, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solar+energy+conversion+efficiencies+in+photosynthesis:+minimizing+the+chlorophyll+antennae+to+maximize+efficiency.">Solar energy conversion efficiencies in photosynthesis: minimizing the chlorophyll antennae to maximize efficiency.</a> Plant Science. 177 (4), 272-280 (2009).</li></ol>

The authors have nothing to disclose.

Within this protocol, focus on the following steps increases the likelihood of generating reproducible, high-quality data. When constructing the reactor stand (step 1), the base must be sturdy with well-alligned vertical supports. Slotted steel has sharp edges, so the addition of safety caps is essential. Bottle platform surfaces need to be completely flat, the magnetic stirrer and bolt heads should both sit below the top layer surface (steps 3.2&#8211;3.6). According to the manufacturer&#39;s instruction, the gas sensor packing liquid should be filled to the &#34;tracing screw for liquid level&#34; for accurate oxygen measurements. This liquid level should be checked regularly as evaporation of the packing liquid can short-circuit the measurement cell. All three gas lines made in step 5.2 should be the same length; this enures that replicates have identical headspace volumes. Before commencing an experiment, it is advisable to test the programmed light regime by logging light intensity over a 24 h period (step 6.11). If increases in liquid temperature are of concern, this test should also include a sealed bottle with an internal temperature probe (step 6.11). When logging, do not exit the data acquisition software window; this will terminate logging. If taking culture samples, be careful not to release headspace gas by opening valves in the wrong sequence (steps 8.2-8.8). When reviewing experimental data be advised that the data acquisition software automatically generates a moving average of the flow rate. This inflates the value of the one or two flow rate readings generated overnight. Curate gas sensor logs manually to rectify this.
The most common setback with this method is the potential to short circuit the gas sensor if the liquid packing level declines. There are two ways this can occur. Firstly, evaporation can slowly reduce the liquid level. However, this is unlikely over a short-term (&#60;7 day) experiment29. Secondly, high respiration rates can pull oxygen into the solution and generate a headspace under-pressure. When light energy is unavailable, microalgae use aerobic respiration to supply the energy required for cellular maintenance and repair28. Hence, in dense cultures during non-illuminated hours, oxygen consumption, and the resulting under-pressure, can be substantial. This sucks packing liquid from the gas sensors into the gas line. The distance the packing liquid travels is proportional to the amount of nighttime respiration. If the packing liquid enters the bottles, this generates an oil slick on the liquid surface.
If high nighttime respiration rates are expected, modifications to the protocol can be made. The simplest way to avoid under-pressure is to leave the bottle headspaces open overnight. This also has the advantage of easing DO levels by decreasing the partial headspace pressure of O2. High DO concentrations are believed to be detrimental to growth as O2 can impede the activity of Rubisco and may trigger oxidative stress30,31. It is not uncommon for culture suspensions to reach 4x oversaturation even when in contact with the atmosphere25,32. To open the headspace, disconnect the gas line from the needle spanning the rubber stopper. Nighttime hours can serve as a window to top up gas sensor packing liquid or manipulate continuous experiments with little impact on data collection. For example, one may alter the culture density, refresh nutrients, add an amendment, or introduce a pathogen. Bottles must be re-sealed, and the gas sensor line reconnected before lights turn back on. The oxygen measurements collected from experiments with closed versus open nighttime headspaces will differ.
When bottles remain sealed, nighttime oxygen consumption reduces the number of moles of O2 in the headspace. This causes packing liquid to creep up the gas sensor line to maintain headspace pressure. When lights turn on, oxygen production resumes. Packing liquid must be pushed back into the gas sensor before flow rate readings commence. This lag is therefore proportional to the degree of nighttime respiration. In this way, when the headspace remains closed, O2 readings represent net O2 production (photosynthetic production - respiratory consumption). Conversely, when the headspace is open at night, atmospheric gas replaces what headspace O2 is consumed, and no packing liquid enters the gas line. The result is that respiratory O2 consumption is not accounted for in O2 production data. This may reduce the accuracy of AFDW biomass growth estimates. However, it should not impact the utility of using daytime O2 production as a metric to compare growth between treatments.
All laboratory PBRs are afflicted by the same limitation; artificial lights cannot replicate the solar spectrum. Microalgae use wavelengths of light between 400&#8211;700 nm for photosynthesis. This region is referred to as photosynthetically active radiation (PAR)33. Sunlight and artificial light vary in their relative contribution of wavelengths within this range. This, alongside favorable temperatures and a constant light supply, means that laboratory growth data often cannot be reliably extrapolated to outdoor conditions. These PBRs can, however, address one of the limitations of laboratory PBR light supply. Sunlight intensity is highly variable throughout the day, with cloud cover generating transient fluctuations in incident PAR. The lighting control software and DMX lighting controller can provide light intensities from 0 to 2400 &#181;molphotons m-2 s-1 and beyond. Light regimes can be broken down into individual increments as short as 1 s. Tunable light intensity allows the user to mimic outdoor light patterns more closely than standard PBR setups. Here, simulated 30 min dawn and dusk intervals fade day and night cycles together (Supplementary Table 1).
Although AFDW density has become the standard measure of growth, this method can require substantial culture volumes, a 2&#8211;3-day processing period, and generates one data point at a time. Further, if conditions become unfavorable and cells die, AFDW density does not discriminate between actively photosynthesizing cells and those that are decomposing. Quantifying the rate of photosynthetic oxygen production serves as an alternative growth proxy. This PBR design can record oxygen production continuously with little user intervention while conserving culture volume. Data resolution could be improved by selecting a gas sensor with a lower measurement cell volume, e.g., 1 mL. Further, if cultures are well-mixed, users may decide to install a spectrophotometer for continuous optical density readings. If temperature control of the medium is desired, a recirculating chiller could be added. These PBRs are a valuable addition to a laboratory looking to expand its microalgal research capacity without heavy financial investment. They are particularly suitable for those working with high alkalinity, high pH species like Spirulina. These PBRs offer light regime flexibility and are valid for rapid, replicated, laboratory growth comparisons.

Microalgae and cyanobacteria, collectively called microalgae for simplicity, are championed for their potential in sustainable biotechnology. They are attractive candidates due to their rapid growth, ability to be cultivated on non-arable land, and for their use of sunlight to drive the conversion of carbon dioxide to biomass1,2,3. Microalgal biomass can be converted into products such as bioenergy in the form of oil or gas, food dyes and nutritional supplements, and materials such as biopolymers1,4,5,6,7. In addition, they can be used to treat wastewater or remediate waterbodies by consuming excess nutrients8,9. Given this, microalgal research is widespread and established. The field grows as society reconsiders the carbon intensity and environmental sustainability of current manufacturing and energy generation approaches.
Three fundamental requirements of laboratory-based microalgal studies are a culture vessel, light source, and method to quantify growth. The term photobioreactor (PBR) describes a set-up in which culture vessels are illuminated10. Commonly, studies of microalgae aim to compare growth between two or more treatments, e.g., different growth media, light regimes, or species11,12,13. For statistical relevance, each condition, e.g., treatment and control, should be replicated. If control and treatment are run simultaneously, this means many PBRs must be monitored and sampled for the duration of an experiment. The challenge with operating multiple PBRs is two-fold. Firstly, supplying a uniform light intensity to each PBR is essential for reproducibility but can be difficult. The amount of light incident on the vessel surface is influenced by its distance from the light source, shading from adjacent vessels, and background light fluctuations14. Secondly, a method to accurately quantify growth must be selected.
Growth is commonly measured by cell count, optical density (OD), chlorophyll A content, dry weight (DW) density, and ash-free dry weight (AFDW) density15. Cell counts, chlorophyll A content, and gravimetric methods are manual processes that produce discrete data points. OD can be measured continuously and non-invasively with a spectrophotometer, provided it is well-calibrated against another method such as AFDW density15. However, OD measurements and Chlorophyll A content can be unreliable as results vary under different culture conditions, e.g., between species and throughout the growth cycle15,16. For Chlorophyll A, the extraction method can also impact pigment yield17. Chlorophyll A content is particularly useful in tracking the growth of microalgae within microbial communities which also contain non-photosynthetic organisms17,18. When choosing a method to determine growth, it is essential to consider the morphology of the suspension. When organisms clump and are not well mixed, OD and cell counts aren&#39;t possible15. A single method is not suited to all experimental applications&#8212;researchers must decide which methods are practical and relevant to their experimental aims.
AFDW is a reliable method that enables growth comparisons among various culture conditions, notably, between species and culture media15,19,20. To calculate AFDW, a sample of microalgal culture is first concentrated, either by filtration or centrifugation, and dried. At this stage, the DW can be determined. Usually, the DW sample contains at least 8&#8211;10% ash-inorganic material such as salts and particulate matter15. DW tracks growth trends but can be skewed if the contribution of inorganics varies. To determine AFDW density, dry biomass is combusted at high temperature; this vaporizes the organic or useful portion while leaving ash (inorganics) behind19. To calculate the AFDW, the weight of the ash fraction is subtracted from that of the DW fraction. Typically, in microalgal suspensions, AFDW ranges from 0.1&#8211;3 g/L12,21,22. Small volumes of dilute suspensions yield little dry biomass, &#60;10 mg. After combustion, ash may only weigh 1 mg. Therefore, depending on the culture density, this method requires volumes between 5&#8211;100 mL and analytical scales accurate to 0.1 mg12,15,19,22. Laboratory PBRs are typically small, a couple of liters at most, hence every liquid sample depletes culture volume. Further, the AFDW method is manual and takes 2&#8211;3 days. For replicated and repetitive experiments, an automated and continuous process is preferable.
For microalgae that use bicarbonate as the carbon source, two additional growth metrics can be measured continuously. Photosynthesis consumes bicarbonate and produces oxygen. Bicarbonate consumption drives up medium pH23. An immersed pH probe can measure this change. Photosynthetic oxygen production increases the medium&#39;s dissolved oxygen (DO) concentration until the medium is saturated. Beyond saturation, oxygen exists as bubbles. Oxygen production is measured by many different techniques: probes gauge DO concentration, manometric devices assess headspace pressure, gas chromatography measures headspace composition, and volumetric sensors record gas outflow24,25,26,27. When oxygen is used as a growth proxy, culture vessels must be fully sealed or allow only gas outflow. For pH and oxygen measurements, carbon must be supplied in the form of bicarbonate, not by CO2 sparging. CO2 sparging decreases the medium pH23 and, as a gas, can disturb oxygen measurements. One advantage of pH and oxygen over optical density is that the method is not compromised if microalgae form clumps. Although indirect, both pH and oxygen are effective in comparing growth between treatments.
PBRs in use today range in complexity. Laboratories may use simple benchtop flasks, custom prototypes, or commercially available products. For research groups seeking to upgrade from flasks, the cost of commercial PBRs or technical skill and part fabrication required to build many prototypes may be a barrier. This manuscript aims to describe the step-by-step design, construction, and operation of laboratory PBRs that bridge this gap. These PBRs have a customizable light regime and monitor growth continuously by recording volumetric oxygen production. This design houses three culture vessels for triplicate replication and can be built with moderate skill and easily accessible materials. This PBR is a valuable addition to a laboratory looking to expand its capacity for microalgal research without investing in very technical or expensive products. When choosing to acquire or build a PBR, researchers must consider the suitability of a design to their culture conditions, financial position, and research questions.

<table><tbody><tr><td>Aluminum channels&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 0.90&amp;rdquo; x 39.37&amp;rdquo;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 2.3 cm x 100 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 4</td><td>LED World</td><td>AC-AR1-1M</td><td>Required as a heat sink</td></tr><tr><td>Bungee cords, small&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 5</td><td>-</td><td>-</td><td>To secure bottles</td></tr><tr><td>Computer - desktop/laptop&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>-</td><td>-</td><td>-</td></tr><tr><td>Data Logger, HOBO U30 USB Weather Station&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>HOBO, Hoskin</td><td>U30-NRC-VIA-10-S100-000</td><td>Records light sensor information</td></tr><tr><td>Digital interface module, Rigamo, 4-channel&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>Ritter</td><td>N/A</td><td>This is to transmit gas sensor data to the computer</td></tr><tr><td>DMX decoder, 12~24 VDC, DMX-CV-4X5A&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>LITECH, LED World</td><td>LT-840-6A</td><td>Transmit messages which alter the light pattern</td></tr><tr><td>DMX lighting controller, SUSHI-RB-RJ&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>Arcolis, Nicolaudie America Inc.</td><td>SUSHI-RB-RJ DMX</td><td>Encodes the lighting program</td></tr><tr><td>Gas sensor packing liquid (Silox)&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1 L</td><td>Ritter</td><td>https://www.ritter.de/en/data-sheets/silox</td><td></td></tr><tr><td>Gas sensor, volumetric&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 3</td><td>Ritter</td><td>MGC-1 V3.4 PMMA (https://www.ritter.de/downloads/mgc-milligascounter-en)</td><td>Measures oxygen production</td></tr><tr><td>Glass bottles, round 1 L with GL45 neck&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 3</td><td>Corning, Capitol Scientific</td><td>1395-1L</td><td>Culture vessels</td></tr><tr><td>Hardware - end caps for slotted steel&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 10</td><td>Paulin, Home Depot</td><td>142-612</td><td>To cover sharp edges of slotted steel</td></tr><tr><td>Hardware - eye hooks&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 6</td><td>-</td><td>-</td><td>To secure bottles</td></tr><tr><td>Hardware - metal corner braces (large)&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 4&quot; x 4&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 10 cm x 10 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 8</td><td>-</td><td>-</td><td>Larger brackets to construct metal stand</td></tr><tr><td>Hardware - metal corner braces (small)&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 2 1/2&quot; x 2 1/2&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 6.4 cm x 6.4 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 6</td><td>-</td><td>-</td><td>Small brackets to connect bottle platforms to PVC pipe</td></tr><tr><td>Hardware - metal corner gussets&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 3&quot; x 3&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 7.6 cm x 7.6 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 6</td><td>Paulin, Home Depot</td><td>142-616</td><td>Flat brackets to construct metal stand</td></tr><tr><td>Hardware - piano hinge&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 36&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 91 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>-</td><td>-</td><td>Connects two halves of PVC pipe</td></tr><tr><td>Hardware - rivets&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 40</td><td>-</td><td>-</td><td>To attach piano hinge to PVC tubing</td></tr><tr><td>Hardware - set of bolts, nuts, washers&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 60</td><td>-</td><td>-</td><td>Long thin bolts are required to secure bottle platforms around magentic stirrers</td></tr><tr><td>Hardware - set of bolts, nuts, washers&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 30</td><td>-</td><td>-</td><td>Larger shorter bolts are required to build the metal stand</td></tr><tr><td>LED driver, constant voltage, 96W 24VDC UL Listed IP65 Driver Class 2 regulated power supply&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>Magnitude Lighting, LED World</td><td>CVN96L24DC</td><td>Regulates power to the lights</td></tr><tr><td>LED lights, Cinco Bright LED Flex Strip&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 4 m roll</td><td>EvenBright, LED World</td><td>FA128M57-4M-24V-X</td><td>Roll is trimmed into 4 x 1 m lengths and secured inside the PVC tube</td></tr><tr><td>Light meter, handheld with submersible sperical probe&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>LI-COR</td><td>LA-250A</td><td>Calibrate the reactors light intensity</td></tr><tr><td>Light sensors Photosynthetic Light (PAR) Smart Sensor&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 2</td><td>HOBO, Hoskin</td><td>S-LIA-M003</td><td>Only one is required however two would be good practice in case one malfunctioned</td></tr><tr><td>Magnetic stirrers (MIXdrive 1 XS) with external control units and power supply (MIXcontrol eco)&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 3</td><td>2Mag, 2MAG USA</td><td>MF 40300</td><td>Stirrers sit sandwiched in bottle platforms</td></tr><tr><td>Metal plate&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 24&quot; x 8&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 61 cm x 20.3 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>-</td><td>-</td><td>This is a surface on which to secure electronics, it is attached to the back of the reactor</td></tr><tr><td>Pipe, white PVC&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 6&quot; diameter x 42&quot; high&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 15.2 cm x 106.7 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>-</td><td>-</td><td>Cut lengthwise in two halves, used to house lights and bottles</td></tr><tr><td>Plastic (HDPE) sheets&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 4&quot; x 4&quot; x 1/4&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 10 cm x 10 cm x 1 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 6</td><td>Inventables</td><td>30291-01</td><td>For bottle platforms which house magentic stirrers</td></tr><tr><td>Rubber stoppers - GL45 size&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 3</td><td>Duran, VWR</td><td>76289-760</td><td>Seals culture vessels</td></tr><tr><td>Screw caps - with aperture and GL45 neck&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 3</td><td>Corning, Capitol Scientific</td><td>1395-45HTSC</td><td>Generates seal of culture vessels</td></tr><tr><td>Slotted angle steel lengths&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 1-1/2&quot; X 48&quot; x 0.074&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 3.8 cm x 122 cm x 0.19 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 6</td><td>Paulin, Home Depot</td><td>142-202</td><td>Makes up the body of the metal stand</td></tr><tr><td>Slotted flat steel lenghts&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 1-3/8&quot; x 48&quot; x 0.074&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 3.5 cm x 122 cm x 0.19 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 3</td><td>Paulin, Home Depot</td><td>142-222</td><td>Makes up the body of the metal stand</td></tr><tr><td>Software - Easy Stand Alone (ESA)</td><td></td><td>https://www.dmxsoft.com/#apps</td><td>AKA LED control software</td></tr><tr><td>Software - Rigamo v3.1</td><td></td><td></td><td>AKA data acquisition software</td></tr><tr><td>Software - Storage Upgrade Tools (SUT)</td><td></td><td>https://store.dmxsoft.com//</td><td></td></tr><tr><td>Stir bar&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: 1&quot; x 5/16&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: 2.5 cm x 0.8 cm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 3</td><td>Fisherbrand</td><td>14-513-59</td><td>Stirs culture</td></tr><tr><td>Switch box&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1</td><td>-</td><td>-</td><td>Turns power on/off to reactor</td></tr><tr><td>Syringe, 10 mL&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: Multiple</td><td>-</td><td>-</td><td>Optional if you wish to extract culture</td></tr><tr><td>Tube adaptor fittings, plastic - Stopcock 1-way&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 6</td><td>Masterflex, Cole Palmer</td><td>RK-12023-33</td><td>Close/open culture vessel line</td></tr><tr><td>Tube adaptor fittings, plastic - variety of male and female luer lock fittings&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: to fit 1/16&quot; and 1/8&quot; ID tubing&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: to fit 1.59 mm and 3.18 mm tubing&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: Multiple packets</td><td>Masterflex, Cole Palmer</td><td>RK-30800-16; RK-30800-18; RK-45518-26; RK-45501-00; RK-45501-04</td><td>Many different combinations can achieve the same end result, best to order a variety of fittings</td></tr><tr><td>Tube adaptor fittings, plastic - variety of straight connectors&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: to fit 1/16&quot; and 1/8&quot; ID tubing&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: to fit 1.59 mm and 3.18 mm tubing&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: Multiple packets</td><td>Masterflex, Cole Palmer</td><td>RK-40616-04</td><td>Many different combinations can achieve the same end result, best to order a variety of fittings</td></tr><tr><td>Tubing, flexible, transparent&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: ID=1/16&quot;, OD=1/8&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: ID = 1.59 mm, 0D = 3.18 mm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 4 m</td><td>Masterflex, Cole Palmer</td><td>RK-06422-02</td><td>Line from culture vessel to gas sensor</td></tr><tr><td>Tubing, flexible, transparent&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: ID=1/8&quot;, OD=1/4&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: ID = 3.18 mm, 0D = 6.35 mm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 2 m</td><td>Masterflex, Cole Palmer</td><td>RK-06422-05</td><td>Gas sensor standard tubing size</td></tr><tr><td>Tubing, rigid, transparent&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: ID=1/16&quot;, OD=1/8&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: ID = 1.59 mm, 0D = 3.18 mm&lt;br/&gt; &lt;strong&gt;Quantiy&lt;/strong&gt;: 1 m</td><td>Masterflex, Cole Palmer</td><td>RK-06605-27</td><td>Spans rubber stopper allowing gas to exit</td></tr><tr><td>Tubing, rigid, transparent&lt;br/&gt; &lt;strong&gt;Imperial&lt;/strong&gt;: ID=1/8&quot;, OD=1/4&quot;&lt;br/&gt; &lt;strong&gt;Metric&lt;/strong&gt;: ID = 3.18 mm, 0D = 6.35 mm&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1 m</td><td>Masterflex, Cole Palmer</td><td>RK-06605-30</td><td>Spans rubber stopper allowing gas to exit</td></tr><tr><td>Zip ties, small&lt;br/&gt; &lt;strong&gt;Quantity&lt;/strong&gt;: 1 packet</td><td></td><td></td><td>Secure tube fittings</td></tr></tbody></table>

operation of laboratory photobioreactors with online growth measurements and customizable light regimes

The laboratory study of microalgae can be experimentally challenging. In addition to the cultivation requirements of non-photosynthetic microorganisms, phototrophs also require illumination. Routinely, researchers seek to provide custom light supplies, i.e., vary the light intensity and time over which it is delivered. Such flexibility is difficult with standard benchtop lights. Usually, cultivation studies also require growth comparisons between experimental treatments. Frequently, growth is assessed over an extended duration, e.g., multiple times a day over a week-long trial. Manual measurements can be time-consuming and lack data resolution. Therefore, photobioreactors (PBRs) with automatic growth monitoring and customizable light supply are useful for replicated experiments with multiple treatments. The current work presents the design, construction, and operation of laboratory PBRs. The materials are easily sourced and relatively inexpensive. The design could be duplicated with moderate skill. Each structure has a footprint of ~40 cm2 and hosts three 1 L glass bottles for triplicate replication. Bottles rest upon platforms containing magnetic stirrers and are arranged vertically within a 1 m high and 15 cm diameter polyvinyl chloride (PVC) pipe. The pipe interior is lined with light-emitting diodes (LEDs). These LEDs produce continuous light intensities from 0&#8211;2400 &#181;mol photons m-2 s-1 of photosynthetically active radiation (PAR). Users design a custom lighting program. The light intensity can be adjusted each second or held constant for longer durations. Oxygen produced from photosynthesis exits each bottle via a one-way volumetric gas sensor. Software is used to record gas sensor data. The amount of oxygen produced can be correlated to biomass growth. If biomass samples are required, a syringe can be used to extract culture. The method is suited for microalgae grown with bicarbonate as the carbon source. These PBRs are valuable to a laboratory that requires replicated experiments, light regime flexibility, and continuous high-resolution growth data.

Watch this Scientific Journal Video about Operation of Laboratory Photobioreactors with Online Growth Measurements and Customizable Light Regimes at JoVE.com

Operation of Laboratory Photobioreactors with Online Growth Measurements and Customizable Light Regimes

This publication describes the design of laboratory photobioreactors (PBRs) with customizable light regimes. The growth of cyanobacteria or microalgae, using bicarbonate as their carbon source, is monitored continuously by measuring volumetric oxygen production. These PBRs facilitate rapid, replicated laboratory growth comparisons with little user intervention during experiments.

The laboratory study of microalgae can be experimentally challenging. In addition to the cultivation requirements of non-photosynthetic microorganisms, ...

operation-laboratory-photobioreactors-with-online-growth-measurements

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

2.1K Views.  University of Calgary. This publication describes the design of laboratory photobioreactors (PBRs) with customizable light regimes. The growth of cyanobacteria or microalgae, using bicarbonate as their carbon source, is monitored continuously by measuring volumetric oxygen production. These PBRs facilitate rapid, replicated laboratory growth comparisons with little user intervention during experiments.

Video: Operation of Laboratory Photobioreactors with Online Growth Measurements and Customizable Light Regimes

Optimize Flue Gas Settings to Promote Microalgae Growth in Photobioreactors via Computer Simulations

Flue gas from power plants can promote algal cultivation and reduce greenhouse gas emissions1. Microalgae not only capture solar energy more efficiently than plants3, but also synthesize advanced biofuels2-4. Generally, atmospheric CO2 is not a sufficient source for supporting maximal algal growth5. On the other hand, the high concentrations of CO2 in industrial exhaust gases have adverse effects on algal physiology. Consequently, both cultivation conditions (such as nutrients and light) and the control of the flue gas flow into the photo-bioreactors are important to develop an efficient &ldquo;flue gas to algae&rdquo; system. Researchers have proposed different photobioreactor configurations4,6 and cultivation strategies7,8 with flue gas. Here, we present a protocol that demonstrates how to use models to predict the microalgal growth in response to flue gas settings. We perform both experimental illustration and model simulations to determine the favorable conditions for algal growth with flue gas. We develop a Monod-based model coupled with mass transfer and light intensity equations to simulate the microalgal growth in a homogenous photo-bioreactor. The model simulation compares algal growth and flue gas consumptions under different flue-gas settings. The model illustrates: 1) how algal growth is influenced by different volumetric mass transfer coefficients of CO2; 2) how we can find optimal CO2 concentration for algal growth via the dynamic optimization approach (DOA); 3) how we can design a rectangular on-off flue gas pulse to promote algal biomass growth and to reduce the usage of flue gas. On the experimental side, we present a protocol for growing Chlorella under the flue gas (generated by natural gas combustion). The experimental results qualitatively validate the model predictions that the high frequency flue gas pulses can significantly improve algal cultivation.

Flue gas from power plants is a cheap CO2 source for algal growth. We have built prototype &#34;flue gas to algal cultivation&#34; systems and described how to scale up the algal cultivation process. We have demonstrated the use of a mass-transfer bio-reaction model to simulate and to design the optimal operation of flue gas for the growth of Chlorella sp. in algal photo-bioreactors.

Flue gas from power plants can promote algal cultivation and reduce greenhouse gas emissions1. Microalgae not only capture solar energy more efficiently ...

Environment

In Situ Measurement and Correlation of Cell Density and Light Emission of Bioluminescent Bacteria

There is a considerable number of bacterial species capable of emitting light. All of them share the same gene cluster, namely the lux operon. Despite this similarity, these bacteria show extreme variations in characteristics like growth behavior, intensity of light emission or regulation of bioluminescence. The method presented here is a newly developed assay that combines recording of cell growth and bioluminescent light emission intensity over time utilizing a plate reader. The resulting growth and light emission characteristics can be linked to important features of the respective bacterial strain, such as quorum sensing regulation. The cultivation of a range of bioluminescent bacteria requires a specific medium (e.g., artificial sea water medium) and defined temperatures. The easy to handle, non-bioluminescent standard-research bacterium Escherichia coli (E. coli), on the other hand, can be cultivated inexpensively in high quantities in laboratory scale. Exploiting E. coli by introducing a plasmid containing the whole lux operon can simplify experimental conditions and additionally opens up many possibilities for future applications. The expression of all lux genes utilizing an E. coli expression strain was achieved by construction of an expression plasmid via Gibson cloning and insertion of four fragments containing seven lux genes and three rib genes of the lux-rib operon into a pET28a vector. E. coli based lux gene expression can be induced and controlled via Isopropyl-&#946;-D-thiogalactopyranosid (IPTG) addition resulting in bioluminescent E. coli cells. The advantages of this system are to avoid quorum sensing regulation restrictions and complex medium compositions along with non-standard growth conditions, such as defined temperatures. This system enables analysis of lux genes and their interplay, by the exclusion of the respective gene from the lux operon, or even addition of novel genes, exchanging the luxAB genes from one bacterial strain by another, or analyzing protein complexes, such as luxCDE.

In Situ Measurement and Correlation of Cell Density and Light Emission of Bioluminescent Bacteria

Bioluminescent bacteria regulate light production through a variety of mechanisms, such as quorum sensing. This novel method allows the in situ investigation of bioluminescence and the correlation of light emission to cell density. An artificial bioluminescent Escherichia coli system allows the characterization of lux operons, Lux proteins, and their interplay.

There is a considerable number of bacterial species capable of emitting light. All of them share the same gene cluster, namely the lux operon. Despite ...

Biochemistry

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 ...

Bioengineering

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.

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 ...

Cultivation of Green Microalgae in Bubble Column Photobioreactors and an Assay for Neutral Lipids

There is significant interest in the study of microalgae for engineering applications such as the production of biofuels, high value products, and for the treatment of wastes. As most new research efforts begin at laboratory scale, there is a need for cost-effective methods for culturing microalgae in a reproducible manner. Here, we communicate an effective approach to culture microalgae in laboratory-scale photobioreactors, and to measure the growth and neutral lipid content of that algae. Instructions are also included on how to set up the photobioreactor system. Although the example organisms are species of Chlorella and Auxenochlorella, this system can be adapted to cultivate a wide range of microalgae, including co-cultures of algae with non-algae species. Stock cultures are first grown in bottles to produce inoculum for the photobioreactor system. Algae inoculum is concentrated and transferred to photobioreactors for cultivation in batch mode. Samples are collected daily for the optical density readings. At the end of the batch culture, cells are harvested by centrifuge, washed, and freeze dried to obtain a final dry weight concentration. The final dry weight concentration is used to create a correlation between the optical density and the dry weight concentration. A modified Folch method is subsequently used to extract total lipids from the freeze-dried biomass and the extract is assayed for its neutral lipid content using a microplate assay. This assay has been published previously but protocol steps were included here to highlight critical steps in the procedure where errors frequently occur. The bioreactor system described here fills a niche between simple flask cultivation and fully-controlled commercial bioreactors. Even with only 3-4 biological replicates per treatment, our approach to culturing algae leads to tight standard deviations in the growth and lipid assays.

Here, we present a protocol to construct lab-scale bubble column photobioreactors and use them to culture microalgae. It also provides a method for the determination of the culture growth rate and neutral lipid content.

There is significant interest in the study of microalgae for engineering applications such as the production of biofuels, high value products, and for the ...

Microalgae Cultivation and Biomass Quantification in a Bench-Scale Photobioreactor with Corrosive Flue Gases

Photobioreactors are illuminated cultivation systems for experiments on phototrophic microorganisms. These systems provide a sterile environment for microalgal cultivation with temperature, pH, and gas composition and flow rate control. At bench-scale, photobioreactors are advantageous to researchers studying microalgal properties, productivity, and growth optimization. At industrial scales, photobioreactors can maintain product purity and improve production efficiency. The video describes the preparation and use of a bench-scale photobioreactor for microalgal cultivation, including the safe use of corrosive gas inputs, and details relevant biomass measurements and biomass productivity calculations. Specifically, the video illustrates microalgal culture storage and preparation for inoculation, photobioreactor assembly and sterilization, biomass concentration measurements, and a logistic model for microalgal biomass productivity with rate calculations including maximum and overall biomass productivities. Additionally, since there is growing interest in experiments to cultivate microalgae using simulated or real waste gas emissions, the video will cover the photobioreactor equipment adaptations necessary to work with corrosive gases and discuss safe sampling in such scenarios.

Bench-scale, axenic cultivation facilitates microalgal characterization and productivity optimization before subsequent process scale-up. Photobioreactors provide the necessary control for reliable and reproducible microalgal experiments and can be adapted to safely cultivate microalgae with the corrosive gases (CO2, SO2, NO2) from municipal or industrial combustion emissions.

Photobioreactors are illuminated cultivation systems for experiments on phototrophic microorganisms. These systems provide a sterile environment for ...

Design and Implementation of an Automated Illuminating, Culturing, and Sampling System for Microbial Optogenetic Applications

Optogenetic systems utilize genetically-encoded proteins that change conformation in response to specific wavelengths of light to alter cellular processes. There is a need for culturing and measuring systems that incorporate programmed illumination and stimulation of optogenetic systems. We present a protocol for building and using a continuous culturing apparatus to illuminate microbial cells with programmed doses of light, and automatically acquire and analyze images of cells in the effluent. The operation of this apparatus as a chemostat allows the growth rate and the cellular environment to be tightly controlled. The effluent of the continuous cell culture is regularly sampled and the cells are imaged by multi-channel microscopy. The culturing, sampling, imaging, and image analysis are fully automated so that dynamic responses in the fluorescence intensity and cellular morphology of cells sampled from the culture effluent are measured over multiple days without user input. We demonstrate the utility of this culturing apparatus by dynamically inducing protein production in a strain of Saccharomyces cerevisiae engineered with an optogenetic system that activates transcription.

We designed a continuous culturing apparatus for use with optogenetic systems to illuminate cultures of microbes and regularly image cells in the effluent with an inverted microscope. The culturing, sampling, imaging, and image analysis are fully automated so that dynamic responses to illumination can be measured over several days.

Optogenetic systems utilize genetically-encoded proteins that change conformation in response to specific wavelengths of light to alter cellular ...

Construction and Setup of a Bench-scale Algal Photosynthetic Bioreactor with Temperature, Light, and pH Monitoring for Kinetic Growth Tests

The optimal design and operation of photosynthetic bioreactors (PBRs) for microalgal cultivation is essential for improving the environmental and economic performance of microalgae-based biofuel production. Models that estimate microalgal growth under different conditions can help to optimize PBR design and operation. To be effective, the growth parameters used in these models must be accurately determined. Algal growth experiments are often constrained by the dynamic nature of the culture environment, and control systems are needed to accurately determine the kinetic parameters. The first step in setting up a controlled batch experiment is live data acquisition and monitoring. This protocol outlines a process for the assembly and operation of a bench-scale photosynthetic bioreactor that can be used to conduct microalgal growth experiments. This protocol describes how to size and assemble a flat-plate, bench-scale PBR from acrylic. It also details how to configure a PBR with continuous pH, light, and temperature monitoring using a data acquisition and control unit, analog sensors, and open-source data acquisition software.

This paper describes the assembly process and operation of a bench-scale photosynthetic bioreactor that can be used, in conjunction with other methods, to estimate pertinent kinetic growth parameters. This system continuously monitors the pH, light, and temperature using sensors, a data acquisition and control unit, and open-source data acquisition software.

The optimal design and operation of photosynthetic bioreactors (PBRs) for microalgal cultivation is essential for improving the environmental and economic ...

Generic Protocol for Optimization of Heterologous Protein Production Using Automated Microbioreactor Technology

A core business in industrial biotechnology using microbial production cell factories is the iterative process of strain engineering and optimization of bioprocess conditions. One important aspect is the improvement of cultivation medium to provide an optimal environment for microbial formation of the product of interest. It is well accepted that the media composition can dramatically influence overall bioprocess performance. Nutrition medium optimization is known to improve recombinant protein production with microbial systems and thus, this is a rewarding step in bioprocess development. However, very often standard media recipes are taken from literature, since tailor-made design of the cultivation medium is a tedious task that demands microbioreactor technology for sufficient cultivation throughput, fast product analytics, as well as support by lab robotics to enable reliability in liquid handling steps. Furthermore, advanced mathematical methods are required for rationally analyzing measurement data and efficiently designing parallel experiments such as to achieve optimal information content.
The generic nature of the presented protocol allows for easy adaption to different lab equipment, other expression hosts, and target proteins of interest, as well as further bioprocess parameters. Moreover, other optimization objectives like protein production rate, specific yield, or product quality can be chosen to fit the scope of other optimization studies. The applied Kriging Toolbox (KriKit) is a general tool for Design of Experiments (DOE) that contributes to improved holistic bioprocess optimization. It also supports multi-objective optimization which can be important in optimizing both upstream and downstream processes.

This manuscript describes a generic approach for tailor-made design of microbial cultivation media. This is enabled by an iterative workflow combining Kriging-based experimental design and microbioreactor technology for sufficient cultivation throughput, which is supported by lab robotics to increase reliability and speed in liquid handling media preparation.

A core business in industrial biotechnology using microbial production cell factories is the iterative process of strain engineering and optimization of ...

Use of High-Throughput Automated Microbioreactor System for Production of Model IgG1 in CHO Cells

Automated microscale bioreactors (15 mL) can be a useful tool for cell culture engineers. They facilitate the simultaneous execution of a wide variety of experimental conditions while minimizing potential process variability. Applications of this approach include: clone screening, temperature and pH shifts,&#160;media and supplement optimization. Furthermore, the small reactor volumes are conducive to large Design of Experiments that investigate a wide range of conditions. This allows upstream processes to be significantly optimized before scale-up where experimentation is more limited in scope due to time and economic constraints. Automated microscale bioreactor systems offer various advantages over traditional small scale cell culture units, such as shake flasks or spinner flasks. However, during pilot scale process development&#160;significant care must be taken to ensure that these advantages are realized. When run with care, the system can enable high level automation, can be programmed to run DOE&#39;s with a higher number of variables&#160;and can reduce sampling time when integrated with a nutrient analyzer or cell counter. Integration of the expert-derived heuristics presented here, with current automated microscale bioreactor experiments can minimize common pitfalls that hinder meaningful results. In the extreme, failure to adhere to the principles laid out here can lead to equipment damage that requires expensive repairs. Furthermore, the microbioreactor systems have small culture volumes&#160;making characterization of cell culture conditions difficult. The number and amount of samples taken in-process in batch mode culture is limited as operating volumes cannot fall below 10 mL. This method will discuss the benefits and drawbacks of microscale bioreactor systems.

A detailed protocol for the concurrent operation of 48 parallel cell cultures under varied conditions in a microbioreactor system is presented. Cell culture process, harvest and subsequent antibody titer analysis are described.

Automated microscale bioreactors (15 mL) can be a useful tool for cell culture engineers. They facilitate the simultaneous execution of a wide variety of ...

Operation of Laboratory Photobioreactors with Online Growth Measurements and Customizable Light Regimes

Summary

Explore More Videos

Chapters in this video

Optimize Flue Gas Settings to Promote Microalgae Growth in Photobioreactors via Computer Simulations

In Situ Measurement and Correlation of Cell Density and Light Emission of Bioluminescent Bacteria

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

Operation of a Benchtop Bioreactor

Cultivation of Green Microalgae in Bubble Column Photobioreactors and an Assay for Neutral Lipids

Microalgae Cultivation and Biomass Quantification in a Bench-Scale Photobioreactor with Corrosive Flue Gases

Design and Implementation of an Automated Illuminating, Culturing, and Sampling System for Microbial Optogenetic Applications

Construction and Setup of a Bench-scale Algal Photosynthetic Bioreactor with Temperature, Light, and pH Monitoring for Kinetic Growth Tests

Generic Protocol for Optimization of Heterologous Protein Production Using Automated Microbioreactor Technology

Use of High-Throughput Automated Microbioreactor System for Production of Model IgG1 in CHO Cells