This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant No. 1546595. Any opinion, findings, and conclusions or recommendations expressed in this material are those of the authors and do not necessarily reflect the views of the National Science Foundation. The work was also supported by a University of Iowa Graduate and Professional Student Government research grant, and the University of Iowa Foundation, Allen S. Henry endowment. Research was conducted in the W. M. Keck Phytotechnologies Laboratory. The authors would like to thank the University of Iowa power plant staff, especially Mark Maxwell, for expertise and financial support for the simulated flue gases. The authors would also like to acknowledge Emily Moore for her assistance with sampling and analysis and Emily Greene for her assistance and participation in the protocol video.

1. Safe use and sampling of a photobioreactor sparged with corrosive gases
NOTE: This method does not describe appropriate procedures for safe sampling of microalgal cultures that produce or consume highly flammable gases.
<ol>
	<li>Manage toxic gas as a risk to human health. 
	NOTE: Per the University of Iowa's Chemical Hygiene Plan, the authors worked with the University Fire Safety Coordinator and University Environmental Health &#38; Safety Industrial Hygiene Officer to develop a safety protocol for working with the toxic gases.</li>
	<li>Set up a toxic gas monitoring system with sensors for each of the toxic gases in use. Calibrate the sensors according to manufacturer instructions. Bump test (check for sensor and alarm functionality with calibration gases) frequently, according to manufacturer instructions. Locate the gas monitor just outside the fume hood.</li>
	<li>Prior to beginning any corrosive gas experiments, notify nearby personnel of the risk and alarm system. Also, notify the appropriate local emergency responders. Post signs on laboratory entrances specifying which hazardous gases are in use.
	<ol>
		<li>Instruct all nearby personnel to evacuate if toxic gas is detected. Notify laboratory supervisors and emergency response personnel. 
		NOTE: In a power outage, the gas-regulating tower will stop gas flow when it loses power. However, if the room heating, ventilation, and air conditioning (HVAC) system or fume hood go down without a power outage, this will result in leaking toxic gas.</li>
	</ol>
	</li>
	<li>Model the possible accumulated concentration of toxic gases in the room if the fume hood were to fail using the American Industrial Hygiene Association's (AIHA) mathematical modeling spreadsheet IH MOD8 for each gas.
	<ol>
		<li>Obtain the room supply/exhaust air rate, Q, in m3 min-1 from building HVAC maintenance personnel or HVAC technician. Calculate the volume, V, of the laboratory (L x W x H) in m3. Calculate the contaminant emission rate, G, of each type of toxic gas in mg min-1, using Equation 1 adapted from the ideal gas law: 
		<img alt="Equation 1" align="center" src="/files/ftp_upload/60566/60566eq1a.jpg" /> (1) 
		where P is the fraction of pressure exerted by the toxic gas at 1 atm (ppm gas/106 ppm), Qgas is the flow rate of the gas in L min-1, R is the universal gas constant (0.082057 L&#183;atm&#183;mol-1&#183;K-1), T is temperature in K, and MW is the gas' molecular weight in g mol-1.</li>
		<li>Use the values for V, Q, and G for each gas (calculated in step 1.4.1) in the "Well-Mixed Room Model With option to cease generation and model room purge" algorithm in the IH MOD spreadsheet to calculate the accumulated room gas concentrations for each gas over a 1440 min (24 h) simulation period. Compare these values to the exposure limits (Table 2)9.</li>
	</ol>
	</li>
</ol>
2. Preparation of the microalgal inoculum
<ol>
	<li>Prepare the Scenedesmus obliquus inoculum, or other microalgal species selected for the photobioreactor, prior to the start of the experiment by transferring it from storage, whether cryopreserved or cultured on agar media.
	<ol>
		<li>Add 30&#8722;50 mL of sterile microalgal growth medium (triple-nitrogen Bold's basal medium [3N-BBM], Table 3) to a sterile (150&#8722;250 mL) shake flask capped with a foam stopper. 
		NOTE: Unless the flask is sparged, only one-fifth of the shake flask volume should be occupied by liquid media.</li>
		<li>Use a biosafety cabinet to maintain sterility when transferring cells to a slant or shake flask. For cultures on agar, use a sterile loop to transfer microalgae from its agar plate or slant to the shake flask. For cryopreserved cultures, gradually thaw the cryopreserved sample and rinse away the cryoprotectant according to the chosen protocol10, then add the cells to the shake flask.</li>
		<li>Culture cells in 3N-BBM at 20&#8722;25 &#176;C under 16 h:8 h light:dark and shaking at 115&#8722;130 rpm.</li>
		<li>Track microalgal growth over time using optical density (OD) measurements (as in sections 5 and 6). Allow the microalgae to reach its exponential growth phase (2&#8722;4 days) before transferring cells to the photobioreactor. 
		NOTE: Depending on the goal of the experiment, the cells may be rinsed of culture medium (this study) and/or concentrated with multiple centrifugation steps prior to inoculation of the bioreactor.</li>
	</ol>
	</li>
</ol>
3. Setup and operation of photobioreactor
<ol>
	<li>Use the photobioreactor (Figure 1) to control temperature, pH, stirring rate, gas flow rates, and input solution flow rates. 
	NOTE: The photobioreactor may be used to control the flow of up to four different input solutions, commonly acid, base, antifoam, and substrate.
	<ol>
		<li>Prepare 100 mL each of 1 N NaOH and 1 N HCl and add each to a 250 mL input solution bottle. Use secondary containment for these solutions.</li>
		<li>Store metered input solutions in autoclavable capped bottles equipped with dip tubes and a vent tube with a sterile inline air filter. Connect the dip tubes to the photobioreactor's four input ports using autoclavable tubing and, during photobioreactor operation, submerse the dip tubes in the input solutions. Pass the 1.6 mm inside dimeter (ID) autoclavable tubing between the input solutions and their ports through separate peristaltic pumps which may be controlled either by manually selected flow rates or by feedback from pH and foam-level probes (in the case of acid, base, and antifoam solutions).</li>
	</ol>
	</li>
	<li>Calibrate the photobioreactor pH meter prior to autoclaving.
	<ol>
		<li>Connect the pH probe to the photobioreactor control tower by fitting the probe to the connecting line and twisting to lock. Use pH 4 and pH 7 buffers to calibrate the pH meter. Wait until values have stabilized before accepting the pH meter value.</li>
		<li>Disconnect the pH probe from the pH meter cord that connects the probe to the control tower.</li>
		<li>Surface sterilize with 70% ethanol or autoclave the probe with the reactor. To autoclave, cap the pH probe tightly with aluminum foil. 
		NOTE: If the probe is autoclaved, there is a risk of corrosion of the probe interior from steam damage. This capping method does not completely guarantee damage prevention.</li>
		<li>Add 10 mM 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES) buffer to the culture medium to better control pH.</li>
	</ol>
	</li>
	<li>Insert and screw closed the cold finger and exhaust condenser on the photobioreactor headplate.</li>
	<li>Insert the inoculation port and screw tightly in place. Add a length of autoclavable tubing to the section of inoculation port above the photobioreactor headplate. Prior to autoclaving the bioreactor, clamp the tubing closed with an autoclavable hose clamp.</li>
	<li>Attach tubing capped with sterile filters to any unused photobioreactor ports.</li>
	<li>Attach acid and base input solutions to the photobioreactor input ports via autoclavable tubing. Add 1.5 L of culture medium.</li>
	<li>Autoclave the reactor and associated input solutions for 30&#8722;45 min at 121 &#176;C according to working volume. 
	NOTE: If the culture medium is adversely affected by autoclaving, add the media after autoclaving under sterile conditions in a laminar flow hood. The protocol can be paused here. 
	CAUTION: Reactor will be hot after removing from autoclave.</li>
	<li>Attach the impeller motor to the impeller shaft and tighten the fitting.</li>
	<li>Arrange LED light panels symmetrically outside the bioreactor according to illumination requirements.
	<ol>
		<li>Prior to autoclaving, measure and record the light intensity with a photometer. Place the illuminance sensor inside the photobioreactor vessel and face the sensor toward the light source.</li>
	</ol>
	</li>
	<li>Connect up to two gas cylinders to supply the simulated coal-fired power plant emissions (Table 1) to the microalgae in the photobioreactor. 
	NOTE: The gas concentrations used in this study approximated those of the University of Iowa power plant.
	<ol>
		<li>Assemble the connections between the gas cylinder, gas regulating tower, and photobioreactor sparging ring. Attach appropriate regulators capable of 20 psi outlet pressure to the gas cylinders. Attach 6 mm ID pressure resistant tubing to the regulator outlet hose barb and secure with a hose clamp. Attach the other end of the pressure resistant tubing to the gas regulating tower gas inlet using a hose barb to 6 mm stem quick connect fitting secured with a hose clamp. Connect 3.2 mm ID tubing to the gas-regulating tower gas outlet using another 6 mm quick connect fitting and connect the other end of the outlet tubing to the sparging ring port at the photobioreactor headplate. 
		NOTE: For a second input gas, repeat step 3.9.1, but use a T-shaped hose barb to consolidate the two input gas lines to a single section of tube connected to the sparging ring port.</li>
		<li>Set the outlet pressure to 20 psi on each gas regulator and use the bioreactor interface to set experimental gas flow rates. 
		NOTE: Calculate and report the volume of air under standard conditions per volume of liquid per minute (vvm); divide the volumetric gas flow rate by the culture volume. Report in units of per miniute&#173;&#173;&#173;.</li>
	</ol>
	</li>
	<li>When sparging with more than one gas cylinder, confirm the supplied CO2 concentration to the photobioreactor with a CO2 sensor.
	<ol>
		<li>Connect a software (e.g., GasLab) compatible CO2 sensor to the USB port of a computer. Download the most recent software that corresponds to the CO2 sensor model. Open the software and input the sensor model, measurement time interval, and duration of measurement data logging.</li>
		<li>Place the CO2 sensor and combined gas flow tube (prior to connecting the tube with the bioreactor) in a 100&#8722;250 mL, capped, vented vessel (external to the bioreactor). 
		NOTE: During the experiment, the headspace CO2 concentrations may be measured from one of the venting tubes on the photobioreactor headplate.</li>
		<li>Start the CO2 concentration measurements on the user interface and wait for the measurements to equilibrate.</li>
		<li>Use the photobioreactor user interface to adjust the gas flow rates from each tank until the desired total flow rate (0.1 L min-1) and CO2 concentration (12%) is achieved.</li>
	</ol>
	</li>
	<li>On the photobioreactor user interface, use the STIRR function to set the impeller rotation rate. Ensure that the mixing rate is rapid enough for the culture medium to assimilate the sparged gas bubbles. 
	NOTE: Certain microalgal species have weak cell structures and will be damaged or ruptured by high shear force.</li>
</ol>
4. Adapting the photobioreactor and experimental setup for toxic gas use
CAUTION: The corrosive gases in real or simulated flue gas are corrosive and toxic. These gases pose serious risk if inhaled. 
NOTE: This method does not describe appropriate materials for safe cultivation of microbes that produce or consume highly flammable gases (i.e., methane, hydrogen, etc.).
<ol>
	<li>Replace brass, plastic, and standard tubing components with corrosion resistant materials.
	<ol>
		<li>Use stainless steel to reliably resist corrosion from the strong acids formed by the reaction between NOx or SOx and water. Replace plastic quick connect fittings at the gas inlets and outlets on the gas-regulating tower with stainless steel quick connect fittings. Use stainless steel regulators for gas cylinders including the outlet hose barb rather than brass.</li>
		<li>Use polytetrafluoroethylene (PTFE) or natural ethyl vinyl acetate (EVA) tubing to resist corrosion from NOx and SOx gases (respectively) on the connections between the gas cylinder to the gas-regulating tower and the gas-regulating tower to the photobioreactor.</li>
	</ol>
	</li>
	<li>After autoclaving, assemble the photobioreactor and gas cylinders within a walk-in fume hood. Place the photobioreactor on a table inside secondary containment and place gas cylinders in free standing cylinder collars or a cylinder rack.</li>
	<li>After initiating gas flow, use a bubble type liquid leak detector to check for gas leaks in all connections between the gas cylinders and bioreactor. Use a wash bottle filled with a 1:100 (v:v) dilution of dish soap:water to cover the connection with a small stream of soap solution. 
	NOTE: Leaks will be indicated by bubbling at the connections.</li>
	<li>When initiating the microalgal experiments, begin sparging then adjust pH before inoculation (as in standard photobioreactor experiments). 
	NOTE: Buffering the culture medium during corrosive gas experiments is highly recommended since the input gases are strongly acidic.</li>
	<li>Inoculate the photobioreactor by aspirating the prepared microalgal inoculum into a sterile syringe, fitting the syringe to the tubing attached to the inoculation port, opening the inoculation tubing clamp, and depressing the syringe.</li>
	<li>Check the gas monitor, gas cylinder pressures, and photobioreactor twice daily (and prior to sampling) for elevated levels of toxic gas or indication of leaks.</li>
	<li>Limit the fume hood sash opening to a width that allows the bioreactor and gas cylinder regulators to be reached. To avoid inhalation exposure risk, ensure that the opening does not expose personnel above the torso region.</li>
	<li>Turn the gas cylinder regulators to the closed position to cease gas flow to the reactor. Close the fume hood sash and allow 5 min for the hood to evacuate the corrosive gases before sampling the photobioreactor culture.</li>
	<li>Sample within the fume hood either by opening a headplate port and using a sterile serological pipet or drawing culture into a syringe through the inoculation/sampling port. Secure the photobioreactor ports prior to opening the gas cylinders and resuming the experiment.</li>
</ol>
5. Measuring microalgal biomass productivity
<ol>
	<li>Use a calibration curve to relate microalgal culture OD750 measurements to dried microalgal biomass concentrations.
	<ol>
		<li>Prepare several (minimum: 4, minimum working volume: 500 mL) flasks with sterile microalgal medium and inoculate with the species of interest (e.g., S. obliquus in this study).</li>
		<li>Measure the culture OD750 until growth is exponential, and promptly sacrificially sample the flasks by filtering known volumes (minimum of 100 mL) of the contents with a 0.45 &#956;m filter membrane of known mass. Use covered aluminum weigh boats or glass vessels to support the biomass and filters as they are dried.</li>
		<li>Mass the biomass and filters after drying for approximately 18&#8722;24 h in an oven between 80&#8722;100 &#176;C. To verify complete drying, measure again after an additional 2&#8722;3 h to determine if the mass has stabilized.</li>
		<li>Subtract the filter mass from the combined biomass and filter mass to calculate the biomass mass.</li>
		<li>Plot the calibration curve as measured OD750 against the biomass concentration (mass of biomass divided by the volume of filtered culture) and fit the data with a linear regression.</li>
	</ol>
	</li>
</ol>
6. Biomass productivity modeling and rate calculations
<ol>
	<li>Calculate experiment biomass concentrations from OD750 measurements using the calibration curve linear regression (determined in section 5).</li>
	<li>Fit batch microalgal growth data from lag to exponential to stationary phase with a logistic regression (Equation 2) in graphing and statistics software (Table of Materials): 
	<img alt="Equation 2" align="center" src="/files/ftp_upload/60566/60566eq2a.jpg" /> (2) 
	where L is the curve's maximum biomass concentration value, k is the relative steepness of the exponential phase (time-1), xo is the time of the sigmoidal curve's midpoint, and x is time.
	<ol>
		<li>Manually enter the logistic equation above. Select Fit a curve with nonlinear regression on the Analysis tab in the software. On the left of the Parameters: Nonlinear Regression box, choose Create new equation under the New dropdown box. Use the default explicit equation as equation type, name the new function, and define the new function as Y = L/(1+ exp(-k*(x-b))), where b represents xo.</li>
	</ol>
	</li>
	<li>Calculate the overall biomass productivity of the microalgal batch by dividing the difference between the final and initial biomass concentrations by the difference between the final and initial times.</li>
	<li>Calculate the maximum biomass productivity of the microalgal batch from the derivative of Equation 2 (Equation 3) at the sigmoid midpoint, when x = xo. 
	<img alt="Equation 3" align="center" src="/files/ftp_upload/60566/60566eq3a.jpg" /> (3)</li>
</ol>

<ol>
	<li>Obom, K. M., Magno, A., Cummings, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Operation+of+a+Benchtop+Bioreactor.">Operation of a Benchtop Bioreactor.</a> Journal of Visualized Experiments. (79), e50582 (2013).</li><li>Cheah, W. Y., Pau Loke, S., Chang, J. -. S., Ling, T., Juan, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biosequestration+of+atmospheric+CO2+and+flue+gas-containing+CO2+by+microalgae.">Biosequestration of atmospheric CO2 and flue gas-containing CO2 by microalgae.</a> Bioresource Technology. 184, 190-201 (2014).</li><li>Xu, L., Weathers, P. J., Xiong, X. -. R., Liu, C. -. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microalgal+bioreactors:+Challenges+and+opportunities.">Microalgal bioreactors: Challenges and opportunities.</a> Engineering in Life Sciences. 9 (3), 178-189 (2009).</li><li>Tsang, Y. 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> Photobioreactors: Advancements, Applications and Research. , (2017).</li><li>Molitor, H. R., Moore, E. J., Schnoor, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maximum+CO2+Utilization+by+Nutritious+Microalgae.">Maximum CO2 Utilization by Nutritious Microalgae.</a> ACS Sustainable Chemistry &amp; Engineering. 7 (10), 9474-9479 (2019).</li><li>Khan, M. I., Shin, J. H., Kim, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+promising+future+of+microalgae:+current+status,+challenges,+and+optimization+of+a+sustainable+and+renewable+industry+for+biofuels,+feed,+and+other+products.">The promising future of microalgae: current status, challenges, and optimization of a sustainable and renewable industry for biofuels, feed, and other products.</a> Microbial Cell Factories. 17 (1), 36 (2018).</li><li>Benemann, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrogen+production+by+microalgae.">Hydrogen production by microalgae.</a> Journal of Applied Phycology. 12 (3), 291-300 (2000).</li><li>. IH MOD Available from: <a target="_blank" href="https://aiha.org/public-resources/consumer-resources/topics-of-interest/ih-apps-tools">https://aiha.org/public-resources/consumer-resources/topics-of-interest/ih-apps-tools</a> (2019)</li><li>Centers for Disease Control and Prevention, Immediately Dangerous To Life or Health (IDLH) Values. The National Institute for Occupational Safety and Health Available from: <a target="_blank" href="https://www.cdc.gov/niosh/idlh/intridl4.html">https://www.cdc.gov/niosh/idlh/intridl4.html</a> (2019)</li><li>Nakanishi, K., Deuchi, K., Kuwano, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreservation+of+four+valuable+strains+of+microalgae,+including+viability+and+characteristics+during+15+of+cryostorage.">Cryopreservation of four valuable strains of microalgae, including viability and characteristics during 15 of cryostorage.</a> Journal of Applied Phycology. 24 (6), 1381-1385 (2012).</li><li>Bischoff, H. W., Bold, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Some soil algae from Enchanted Rock and related algal species. , (1963).</li><li>Mata, T. M., Martins, A. A., Caetano, N. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microalgae+for+biodiesel+production+and+other+applications:+A+review.">Microalgae for biodiesel production and other applications: A review.</a> Renewable and Sustainable Energy Reviews. 14 (1), 217-232 (2010).</li><li>Wood, A. M., Everroad, R. C., Wingard, L. M., Andersen, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+growth+rates+in+microalgal+cultures.">Measuring growth rates in microalgal cultures.</a> Algal Culturing Techniques. , 270-272 (2005).</li></ol>

The authors have nothing to disclose.

Batch, axenic photobioreactor experiments with regulated pH, temperature, gas flow rate, and gas concentration promote meaningful results by eliminating contamination by non-target algal strains and variability in culture conditions. Accurate pure culture growth kinetics can be obtained even in the presence of corrosive gases (CO2, SO2, NO2), which serve as nutrients, turning waste gases into a valuable product such as animal feed.
Prior to beginning any microa...

Photobioreactors are useful for controlled experiments and cultivation of purer microalgal products than can be achieved by open ponds. Microalgal cultivation in bench-scale photobioreactors supports the development of fundamental knowledge that may be used for process scale-up. Slight changes to environmental conditions can significantly alter microbiological experiments and confound the results1. A sterile process with temperature, pH, and gas sparging control is advantageous for studying microalgal properties and performance under varied conditions. Additionally, the control over input gas concentrations, temperature, shear force from mixing, and medium pH can support diverse species that are otherwise challenging to cultivate. Photobioreactors can be run as a batch process with continuous gas feed and sparging, or as a chemostat flow-through system with continuous gas feed and sparging plus influent and effluent wastewater nutrient inputs. Here, we demonstrate the batch process with continuous gas feed and sparging.
The use of photobioreactors addresses several microalgal cultivation and production challenges. The field generally struggles with concerns of contamination by other microorganisms, efficient substrate utilization (which is especially important in the case of CO2 mitigation or wastewater treatment)2, pH control, illumination variability, and biomass productivity3. Photobioreactors enable researchers to study a wide range of phototrophs in closely-controlled batch systems, where even slow growing species are protected from predators or competing microorganisms4. These batch systems are also better at facilitating greater CO2 utilization rates and biomass productivity because they are closed systems that are more likely to be in equilibrium with supplied gases. Photobioreactor technology also offers pH control, the lack of which has hindered high biomass productivity in past studies5. At bench scale, the level of control offered by photobioreactors is advantageous to researchers. At larger industrial scales, photobioreactors can be used to maintain commercial bioproduct purity and improve production efficiency for nutraceutical, cosmetic, food, or feed applications6.
Microalgae are of great interest for biosequestration of CO2 because they can rapidly fix CO2 as biomass carbon. However, most anthropogenic sources of CO2 are contaminated with other corrosive and toxic gases or contaminants (NOx, SOx, CO, Hg), depending on combustion process fuel source. Growing interest in sustainable CO2 sequestration has prompted development of photobioreactor technologies to treat CO2-rich emissions, such as those from coal-fired power plants (Table 1). Unfortunately, there is inherent risk of human and environmental exposure to the corrosive and toxic contaminants during research and scale-up processes. As such, describing the safe assembly and operation of bioreactors using corrosive gases is necessary and instructive.
This method is for the use of a 2 L bench-scale photobioreactor for the growth of microalgae under carefully controlled experimental conditions. The protocol describes microalgal storage, inoculum preparation, and photobioreactor setup and sterilization. Beyond basic operation, this work describes microalgal biomass measurements and biomass productivity calculations, and adaptation of the equipment for microalgal cultivation with corrosive gases. The protocol described below is appropriate for researchers seeking to exert greater experimental control, optimize microalgal growth conditions, or axenically culture a range of phototrophic microbes. This method does not describe appropriate materials for cultivation of microbes that produce or consume flammable gases (e.g. CH4, H2, etc.)7.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Biostat A bioreactor</td>
			<td>Sartorius Stedim</td>
			<td></td>
			<td>2-liter bioreactor for microbial fermentation; designed to be autoclaved; pH, temperature, gas flow rate control</td>
		</tr>
		<tr>
			<td>Bump test NO2 gas</td>
			<td>Grainger</td>
			<td>GAS34L-112-5</td>
			<td>Calibration gas for MultiRAE gas detector</td>
		</tr>
		<tr>
			<td>Bump test O2, CO, LEL gas</td>
			<td>Grainger</td>
			<td>GAS44ES-301A</td>
			<td>Calibration gas for MultiRAE gas detector</td>
		</tr>
		<tr>
			<td>Bump test SO2 gas</td>
			<td>Grainger</td>
			<td>GAS34L-175-5</td>
			<td>Calibration gas for MultiRAE gas detector</td>
		</tr>
		<tr>
			<td>Corrosion resistant tubing for NO2 gas</td>
			<td>Swagelok</td>
			<td>SS-XT4TA4TA4-6</td>
			<td>PTFE Core Hose Smooth Bore X Series&#8212;Fiber Braid and 304 SS Braid Reinforcement</td>
		</tr>
		<tr>
			<td>Corrosion resistant tubing for SO2 gas</td>
			<td>QC Supply</td>
			<td>120325</td>
			<td>Reinforced Braided Natural EVA Tubing - 1/4&#34; ID</td>
		</tr>
		<tr>
			<td>cozIR 100% CO2 meter</td>
			<td>Gas Sensing Solutions Ltd.</td>
			<td>CM-0121 at CO2meter.com</td>
			<td>CO2 meter for concentrations up to 100%</td>
		</tr>
		<tr>
			<td>cozIR 20% CO2 meter</td>
			<td>Gas Sensing Solutions Ltd.</td>
			<td>CM-0123 at CO2meter.com</td>
			<td>CO2 meter for concentrations up to 20%</td>
		</tr>
		<tr>
			<td>Durapore Membrane Filter, 0.45 &#956;m</td>
			<td>Millipore Sigma</td>
			<td>HVLP04700</td>
			<td>Hydrophilic, plain white, 47 mm diameter, 0.45 &#956;m pore size, PVFD membrane filters</td>
		</tr>
		<tr>
			<td>Gas cylinder regulators</td>
			<td>Praxair</td>
			<td>PRS 40221331-660</td>
			<td>Single-stage stainless steel regulator configured for 0-15 psi outlet assembly diaphragm valve with 1/4&#34; MNPT threads, Stainless steel to resist corrosion from NOx and SOx</td>
		</tr>
		<tr>
			<td>Gas cylinders</td>
			<td>Praxair</td>
			<td>Ulta-zero air, high purity CO2, or custom gas composition</td>
			<td>Dependent on study objectives</td>
		</tr>
		<tr>
			<td>Gas monitoring and leak detection system</td>
			<td>RAE Systems by Honeywell</td>
			<td>MAB3000235E020</td>
			<td>Pumped model that detects O2, SO2, NO2, CO, and LEL</td>
		</tr>
		<tr>
			<td>GasLab software</td>
			<td>GasLab</td>
			<td>v2.0.8.14</td>
			<td>Software for CO2 meter measurements and data logging</td>
		</tr>
		<tr>
			<td>Hose barb</td>
			<td>Grainger</td>
			<td>Item # 3DTN3</td>
			<td>Used to adapt regulators to tubing, Stainless steel to resist corrosion from NOx and SOx</td>
		</tr>
		<tr>
			<td>K30 1% CO2 meter</td>
			<td>Senseair</td>
			<td>CM-0024 at CO2meter.com</td>
			<td>CO2 meter for concentrations less than 1%</td>
		</tr>
		<tr>
			<td>LED grow panels</td>
			<td>Roleadro</td>
			<td>HY-MD-D169-S</td>
			<td>Red &#38; blue LED light panels</td>
		</tr>
		<tr>
			<td>Memosens dissolved oxygen probe</td>
			<td>Endress+ Hauser</td>
			<td>COS22D-19M6/0</td>
			<td>Autoclavable (with precautions) dissolved oxygen probe for bioreactor</td>
		</tr>
		<tr>
			<td>Memosens pH probe</td>
			<td>Endress+ Hauser</td>
			<td>CPS71D-7TB41</td>
			<td>Autoclavable (with precautions) pH probe for bioreactor</td>
		</tr>
		<tr>
			<td>Oven, Isotemp 500 Series</td>
			<td>Fisher Scientific</td>
			<td>13246516GAQ</td>
			<td>Small oven for drying</td>
		</tr>
		<tr>
			<td>Prism GraphPad software</td>
			<td>GraphPad Software</td>
			<td>Version 7.03 or 8.0.1</td>
			<td>Graphing software for data organization, data analysis, and publication-quality graphs</td>
		</tr>
		<tr>
			<td>Stem to hose barb fitting</td>
			<td>Swagelok</td>
			<td>SS-4-HC-A-6MTA</td>
			<td>Stainless Steel Hose Connector, 6 mm Tube Adapter, 1/4 in. Hose ID</td>
		</tr>
		<tr>
			<td>Tubing, dilute acid/base transfer</td>
			<td>Allied Electronics and Automation</td>
			<td>6678441</td>
			<td>Silicone TP Process Tubing; 1.6mm Bore Size; 3000mm Long; Food Grade</td>
		</tr>
		<tr>
			<td>Tubing, gas transfer</td>
			<td>Allied Electronics and Automation</td>
			<td>6678444</td>
			<td>Silicone TP Process Tubing; 3.2mm Bore Size; 3000mm Long; Food Grade</td>
		</tr>
	</tbody>
</table>

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.

A calibration curve for the green microalgae, S. obliquus, harvested in the exponential phase, was established with OD750 and dried biomass concentrations (Figure 2). The linear regression had an R2 value of 0.9996.
An S. obliquus culture was started in a 250 mL Erlenmeyer flask from a culture stored on a refrigerated agar plate. The microalga was inoculated in 3N-BBM with 10 mM HEPES buffer and sparged with 2.2% CO2</s...

Watch this Scientific Journal Video about Microalgae Cultivation and Biomass Quantification in a Bench-Scale Photobioreactor with Corrosive Flue Gases at JoVE.com

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

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 ...

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10.1K Views.  University of Iowa. 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.

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

Analysis of Fatty Acid Content and Composition in Microalgae

A method to determine the content and composition of total fatty acids present in microalgae is described. Fatty acids are a major constituent of microalgal biomass. These fatty acids can be present in different acyl-lipid classes. Especially the fatty acids present in triacylglycerol (TAG) are of commercial interest, because they can be used for production of transportation fuels, bulk chemicals, nutraceuticals (&omega;-3 fatty acids), and food commodities. To develop commercial applications, reliable analytical methods for quantification of fatty acid content and composition are needed. Microalgae are single cells surrounded by a rigid cell wall. A fatty acid analysis method should provide sufficient cell disruption to liberate all acyl lipids and the extraction procedure used should be able to extract all acyl lipid classes.
With the method presented here all fatty acids present in microalgae can be accurately and reproducibly identified and quantified using small amounts of sample (5 mg) independent of their chain length, degree of unsaturation, or the lipid class they are part of.
This method does not provide information about the relative abundance of different lipid classes, but can be extended to separate lipid classes from each other.
The method is based on a sequence of mechanical cell disruption, solvent based lipid extraction, transesterification of fatty acids to fatty acid methyl esters (FAMEs), and quantification and identification of FAMEs using gas chromatography (GC-FID). A TAG internal standard (tripentadecanoin) is added prior to the analytical procedure to correct for losses during extraction and incomplete transesterification.

A method for the determination of fatty acid content and composition in microalgae based on mechanical cell disruption, solvent based lipid extraction, transesterification, and quantification and identification of fatty acids using gas chromatography is described. A tripentadecanoin internal standard is used to compensate for the possible losses during extraction and incomplete transesterification.

A method to determine the content and composition of total fatty acids present in microalgae is described. Fatty acids are a major constituent of ...

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 ...

Quantification of Heavy Metals and Other Inorganic Contaminants on the Productivity of Microalgae

Increasing demand for renewable fuels has researchers investigating the feasibility of alternative feedstocks, such as microalgae. Inherent advantages include high potential yield, use of non-arable land and integration with waste streams. The nutrient requirements of a large-scale microalgae production system will require the coupling of cultivation systems with industrial waste resources, such as carbon dioxide from flue gas and nutrients from wastewater. Inorganic contaminants present in these wastes can potentially lead to bioaccumulation in microalgal biomass negatively impact productivity and limiting end use. This study focuses on the experimental evaluation of the impact and the fate of 14 inorganic contaminants (As, Cd, Co, Cr, Cu, Hg, Mn, Ni, Pb, Sb, Se, Sn, V and Zn) on Nannochloropsis salina growth. Microalgae were cultivated in photobioreactors illuminated at 984 &#181;mol m-2 sec-1 and maintained at pH 7 in a growth media polluted with inorganic contaminants at levels expected based on the composition found in commercial coal flue gas systems. Contaminants present in the biomass and the medium at the end of a 7 day growth period were analytically quantified through cold vapor atomic absorption spectrometry for Hg and through inductively coupled plasma mass spectrometry for As, Cd, Co, Cr, Cu, Mn, Ni, Pb, Sb, Se, Sn, V and Zn. Results show N. salina is a sensitive strain to the multi-metal environment with a statistical decrease in biomass yieldwith the introduction of these contaminants. The techniques presented here are adequate for quantifying algal growth and determining the fate of inorganic contaminants.

Integration of microalgal cultivation with industrial flue gas will ultimately introduce heavy metals and other inorganic compounds into the growth media. This study presents a procedure used to determine the end fate and impact of heavy metals and inorganic contaminants on the growth of Nannochloropsis salina grown in photobioreactors.

Increasing demand for renewable fuels has researchers investigating the feasibility of alternative feedstocks, such as microalgae. Inherent advantages ...

High-throughput Screening of Recalcitrance Variations in Lignocellulosic Biomass: Total Lignin, Lignin Monomers, and Enzymatic Sugar Release

The conversion of lignocellulosic biomass to fuels, chemicals, and other commodities has been explored as one possible pathway toward reductions in the use of non-renewable energy sources. In order to identify which plants, out of a diverse pool, have the desired chemical traits for downstream applications, attributes, such as cellulose and lignin content, or monomeric sugar release following an enzymatic saccharification, must be compared. The experimental and data analysis protocols of the standard methods of analysis can be time-consuming, thereby limiting the number of samples that can be measured. High-throughput (HTP) methods alleviate the shortcomings of the standard methods, and permit the rapid screening of available samples to isolate those possessing the desired traits. This study illustrates the HTP sugar release and pyrolysis-molecular beam mass spectrometry pipelines employed at the National Renewable Energy Lab. These pipelines have enabled the efficient assessment of thousands of plants while decreasing experimental time and costs through reductions in labor and consumables.

Plant cell wall structure and chemistry traits are evaluated to identify ideal feedstocks for biofuels and bio-materials. Standard methods have limitations when applied to large data sets. These high-throughput pretreatment, enzyme saccharification, and pyrolysis-molecular beam mass spectrometry methods compare large numbers of biomass samples with decreased experimental time and cost.

The conversion of lignocellulosic biomass to fuels, chemicals, and other commodities has been explored as one possible pathway toward reductions in the ...

Experimental Protocol for Biodiesel Production with Isolation of Alkenones as Coproducts from Commercial Isochrysis Algal Biomass

The need to replace petroleum fuels with alternatives from renewable and more environmentally sustainable sources is of growing importance. Biomass-derived biofuels have gained considerable attention in this regard, however first generation biofuels from edible crops like corn ethanol or soybean biodiesel have generally fallen out of favor. There is thus great interest in the development of methods for the production of liquid fuels from domestic and superior non-edible sources. Here we describe a detailed procedure for the production of a purified biodiesel from the marine microalgae Isochrysis. Additionally, a unique suite of lipids known as polyunsaturated long-chain alkenones are isolated in parallel as potentially valuable coproducts to offset the cost of biodiesel production. Multi-kilogram quantities of Isochrysis are purchased from two commercial sources, one as a wet paste (80% water) that is first dried prior to processing, and the other a dry milled powder (95% dry). Lipids are extracted with hexanes in a Soxhlet apparatus to produce an algal oil (&#34;hexane algal oil&#34;) containing both traditional fats (i.e., triglycerides, 46-60% w/w) and alkenones (16-25% w/w). Saponification of the triglycerides in the algal oil allows for separation of the resulting free fatty acids (FFAs) from alkenone-containing neutral lipids. FFAs are then converted to biodiesel (i.e., fatty acid methyl esters, FAMEs) by acid-catalyzed esterification while alkenones are isolated and purified from the neutral lipids by crystallization. We demonstrate that biodiesel from both commercial Isochrysis biomasses have similar but not identical FAME profiles, characterized by elevated polyunsaturated fatty acid contents (approximately 40% w/w). Yields of biodiesel were consistently higher when starting from the Isochrysis wet paste (12% w/w vs. 7% w/w), which can be traced to lower amounts of hexane algal oil obtained from the powdered Isochrysis product.

Experimental Protocol for Biodiesel Production with Isolation of Alkenones as Coproducts from Commercial Isochrysis Algal Biomass

Detailed methods are presented for the production of biodiesel along with the co-isolation of alkenones as valuable coproducts from commercial Isochrysis microalgae.

The need to replace petroleum fuels with alternatives from renewable and more environmentally sustainable sources is of growing importance. ...

Pretreatment of Lignocellulosic Biomass with Low-cost Ionic Liquids

A number of ionic liquids (ILs) with economically attractive production costs have recently received growing interest as media for the delignification of a variety of lignocellulosic feedstocks. Here we demonstrate the use of these low-cost protic ILs in the deconstruction of lignocellulosic biomass (Ionosolv pretreatment), yielding cellulose and a purified lignin. In the most generic process, the protic ionic liquid is synthesized by accurate combination of aqueous acid and amine base. The water content is adjusted subsequently. For the delignification, the biomass is placed into a vessel with IL solution at elevated temperatures to dissolve the lignin and hemicellulose, leaving a cellulose-rich pulp ready for saccharification (hydrolysis to fermentable sugars). The lignin is later precipitated from the IL by the addition of water and recovered as a solid. The removal of the added water regenerates the ionic liquid, which can be reused multiple times. This protocol is useful to investigate the significant potential of protic ILs for use in commercial biomass pretreatment/lignin fractionation for producing biofuels or renewable chemicals and materials.

The pretreatment of lignocellulosic biomass with protic low-cost ionic liquids is shown, resulting in a delignified cellulose-rich pulp and a purified lignin. The pulp gives rise to high glucose yields after enzymatic saccharification.

A number of ionic liquids (ILs) with economically attractive production costs have recently received growing interest as media for the delignification of ...

Ammonia Fiber Expansion (AFEX) Pretreatment of Lignocellulosic Biomass

Lignocellulosic materials are plant-derived feedstocks, such as crop residues (e.g., corn stover, rice straw, and sugar cane bagasse) and purpose-grown energy crops (e.g., miscanthus, and switchgrass) that are available in large quantities to produce biofuels, biochemicals, and animal feed. Plant polysaccharides (i.e., cellulose, hemicellulose, and pectin) embedded within cell walls are highly recalcitrant towards conversion into useful products. Ammonia fiber expansion (AFEX) is a thermochemical pretreatment that increases accessibility of polysaccharides to enzymes for hydrolysis into fermentable sugars. These released sugars can be converted into fuels and chemicals in a biorefinery. Here, we describe a laboratory-scale batch AFEX process to produce pretreated biomass on the gram-scale without any ammonia recycling. The laboratory-scale process can be used to identify optimal pretreatment conditions (e.g., ammonia loading, water loading, biomass loading, temperature, pressure, residence time, etc.) and generates sufficient quantities of pretreated samples for detailed physicochemical characterization and enzymatic/microbial analysis. The yield of fermentable sugars from enzymatic hydrolysis of corn stover pretreated using the laboratory-scale AFEX process is comparable to pilot-scale AFEX process under similar pretreatment conditions. This paper is intended to provide a detailed standard operating procedure for the safe and consistent operation of laboratory-scale reactors for performing AFEX pretreatment of lignocellulosic biomass.

Ammonia Fiber Expansion AFEX Pretreatment of Lignocellulosic Biomass

Ammonia fiber expansion (AFEX) is a thermochemical pretreatment technology that can convert lignocellulosic biomass (e.g., corn stover, rice straw, and sugarcane bagasse) into a highly digestible feedstock for both biofuels and animal feed applications. Here, we describe a laboratory-scale method for conducting AFEX pretreatment on lignocellulosic biomass.

Lignocellulosic materials are plant-derived feedstocks, such as crop residues (e.g., corn stover, rice straw, and sugar cane bagasse) and purpose-grown ...

A Novel Method for the Pentosan Analysis Present in Jute Biomass and Its Conversion into Sugar Monomers Using Acidic Ionic Liquid

Recently, ionic liquids (ILs) are used for biomass valorization into valuable chemicals because of their remarkable properties such as thermal stability, lower vapor pressure, non-flammability, higher heat capacity, and tunable solubility and acidity. Here, we demonstrate a method for the synthesis of C5 sugars (xylose and arabinose) from the pentosan present in jute biomass in a one-pot process by utilizing a catalytic amount of Br&#248;nsted acidic 1-methyl-3-(3-sulfopropyl)-imidazolium hydrogen sulfate IL. The acidic IL is synthesized in the lab and characterized using NMR spectroscopic techniques for understanding its purity. The various properties of BAIL are measured such as acid strength, thermal and hydrothermal stability, which showed that the catalyst is stable at a higher temperature (250 &#176;C) and possesses very high acid strength (Ho 1.57). The acidic IL converts over 90% of pentosan into sugars and furfural. Hence, the presenting method in this study can also be employed for the evaluation of pentosan concentration in other kinds of lignocellulosic biomass.

We present a protocol for the synthesis of C5 sugars (xylose and arabinose) from a renewable non-edible lignocellulosic biomass (i.e., jute) with the presence of Br&#248;nsted acidic ionic liquids (BAILs) as the catalyst in water. The BAILs catalyst exhibited better catalytic performance than conventional mineral acid catalysts (H2SO4 and HCl).

Recently, ionic liquids (ILs) are used for biomass valorization into valuable chemicals because of their remarkable properties such as thermal stability, ...

Preparation of Biomass-based Mesoporous Carbon with Higher Nitrogen-/Oxygen-chelating Adsorption for Cu(II) Through Microwave Pre-Pyrolysis

An environment-friendly technique for synthesizing biomass-based mesoporous activated carbon with high nitrogen-/oxygen-chelating adsorption for Cu(II) is proposed. Bagasse impregnated with phosphoric acid is utilized as the precursor. To pyrolyze the precursor, two separate heating modes are used: microwave pyrolysis and conventional electric-heating pyrolysis. The resulting bagasse-derived carbon samples are modified with nitrification and reduction modification. Nitrogen (N)/oxygen (O) functional groups are simultaneously introduced to the surface of activated carbon, enhancing its adsorption of Cu(II) by complexing and ion-exchange. Characterization and copper adsorption experiments are performed to investigate the physicochemical properties of four prepared carbon samples and determine which heating method favors the subsequent modification for doping of N/O functional groups. In this technique, based on analyzing data of nitrogen adsorption, Fourier transform infrared spectroscopy, and batch adsorption experiments, it is proven that microwave-pyrolyzed carbon has more defect sites and, therefore, time-saving effective microwave pyrolysis contributes more N/O species to the carbon, although it leads to a lower specific surface area. This technique offers a promising route to synthesis adsorbents with higher nitrogen and oxygen content and a higher adsorption capacity of heavy-metal ions in wastewater remediation applications.

Preparation of Biomass-based Mesoporous Carbon with Higher Nitrogen-/Oxygen-chelating Adsorption for CuII Through Microwave Pre-Pyrolysis

Here, we present a protocol to synthesize nitrogen/oxygen dual-doped mesoporous carbon from biomass by chemical activation in different pyrolysis modes followed by modification. We demonstrate that the microwave pyrolysis benefits the subsequent modification process to simultaneously introduce more nitrogen and oxygen functional groups on the carbon.

An environment-friendly technique for synthesizing biomass-based mesoporous activated carbon with high nitrogen-/oxygen-chelating adsorption for Cu(II) is ...

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

Summary

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Chapters in this video

Analysis of Fatty Acid Content and Composition in Microalgae

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

Quantification of Heavy Metals and Other Inorganic Contaminants on the Productivity of Microalgae

High-throughput Screening of Recalcitrance Variations in Lignocellulosic Biomass: Total Lignin, Lignin Monomers, and Enzymatic Sugar Release

Experimental Protocol for Biodiesel Production with Isolation of Alkenones as Coproducts from Commercial Isochrysis Algal Biomass

Pretreatment of Lignocellulosic Biomass with Low-cost Ionic Liquids

Ammonia Fiber Expansion (AFEX) Pretreatment of Lignocellulosic Biomass

A Novel Method for the Pentosan Analysis Present in Jute Biomass and Its Conversion into Sugar Monomers Using Acidic Ionic Liquid

Preparation of Biomass-based Mesoporous Carbon with Higher Nitrogen-/Oxygen-chelating Adsorption for Cu(II) Through Microwave Pre-Pyrolysis

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