Name: cultivation of green microalgae in bubble column photobioreactors and an assay for neutral lipids
Uploaded: 2019-01-07T14:00:00.000+00:00
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Description: Watch this Scientific Journal Video about Cultivation of Green Microalgae in Bubble Column Photobioreactors and an Assay for Neutral Lipids at JoVE.com

Support for this research was provided by USDA National Institute of Food and Agriculture Hatch Project ALA0HIGGINS and the Auburn University Offices of the Provost, the Vice President for Research, and the Samuel Ginn College of Engineering. Support was also provided by NSF grant CBET-1438211.

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The authors have nothing to disclose.

The most important consideration when culturing algae is an understanding of the specific needs of the organism or group of organisms. The algae cultivation system described here can be used to culture a wide range of algae but the specific abiotic factors (temperature, media, pH, light intensity, CO2 level, aeration rate) need to be adjusted to the needs of the organism. Note the parameters described here were used for the cultivation of Chlorella and Auxenochlorella. These organisms are of ...

The application of microalgae in engineering and biotechnology has attracted great interest in recent years. Microalgae are being studied for use in wastewater treatment1,2,3,4, biofuel production5,6,7,8, and the production of nutraceuticals and other high value products9,10. Algae are also being genetically modified at greater rates in an effort to improve their fitness for specific engineering applications11,12. Consequently, there is great interest in experimentation with industrially relevant organisms in controlled settings. The purpose of this method is to communicate an effective approach to culture microalgae in a controlled laboratory environment, and to measure the growth and neutral lipid content of that algae. Improving growth rates and neutral lipid content of microalgae have been identified as two key bottlenecks toward commercialization of algal biofuels13.
A wide range of approaches have been used to culture algae for experimental purposes. In general, these approaches can be divided between large-scale outdoor cultivation and small-scale indoor cultivation. Outdoor cultivation in photobioreactors and open ponds is appropriate for experimentation aimed at scaling up processes that have already been proven at laboratory scale (e.g., to test scale-up of a new high-lipid strain of algae)14. However, indoor small-scale cultivation is appropriate when developing new or improved algae strains or performing experiments aimed at understanding biological mechanisms. In these latter cases, a high-degree of experimental control is required to tease out subtle changes in biological behavior. To that end, axenic cultures are often required in order to minimize the complex biotic factors associated with other organisms (e.g. bacteria, other algae) that inevitably grow in large-scale outdoor systems. Even when studying interactions among algae and other organisms, we have found that use of highly-controlled experimental conditions is helpful when examining molecular exchange among organisms15,16,17.
Within the category of small-scale indoor algae cultivation, a range of approaches have been used. Perhaps the most common approach is to grow algae in Erlenmeyer flasks on a shaker table beneath a light bank18,19. Exchange of oxygen and CO2 takes place by passive diffusion through a foam plug in the top of the flask. Some researchers have improved this set-up through active aeration of the flasks20. Another approach is to cultivate algae in bottles, mixed by stir bar and active aeration. Despite their simplicity, we have found that the use of flasks and bottles often leads to inconsistent results among biological replicates. Presumably this is due to position effects - different positions receive different amounts of light, which also affect internal reactor temperatures. Daily rotation of reactors to new positions can help but does not alleviate the problem because certain stages of algae growth (e.g., early exponential) are more sensitive to positional effects than others (e.g., log phase).
On the opposite side of the spectrum of technological sophistication are fully-controlled commercial photobioreactors. These systems continuously monitor and adjust conditions in the reactor to optimize algae growth. They have programmable lighting, real-time temperature control, and pH control. Unfortunately, they are expensive and typically cost several thousand dollars per reactor. Most scientific and engineering journals require biological replication of results, necessitating the purchase of multiple bioreactors. We present here a bubble column reactor system that bridges the divide between the simple (flask) and sophisticated (fully-controlled bioreactor) approaches for lab-scale algae cultivation. Bubble columns use rising gas bubbles to facilitate gas exchange and mix the reactor. This approach provides some degree of control over the lighting and temperature but does so in a way that is cost-effective. Moreover, we have found this system to yield highly consistent results among biological replicates, reducing the required number of biological replicates needed in order to obtain statistically significant results when compared to the flask or bottle approach. We have also used this system to successfully cultivate mixtures of algae and bacteria21. In addition to algae cultivation, we outline a procedure for measuring the neutral lipid content in the cultured algae. The latter method has been published elsewhere22, but we include the procedure here to provide step-by-step instructions for how to employ it successfully.

<table><tbody><tr><td>&lt;strong&gt;Supplies for airlift photobioreactor setup&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1 L Pyrex bottles</td><td>Corning</td><td>16157-191</td><td>For bottle reactors, humidifiers</td></tr><tr><td>1/2&quot; hose clamp</td><td>Home Depot</td><td>UC953A</td><td>or equivalent</td></tr><tr><td>1/4&quot; female luer to barb</td><td>Nordson biomedical</td><td>Nordson FTLL360-6005</td><td>1/4&quot; ID, PP</td></tr><tr><td>1/4&quot; ID, 3/8&quot; OD autoclaveable PVC tubing</td><td>Thermo-Nalgene</td><td>63013-244</td><td>50&#039;</td></tr><tr><td>1/4&quot; in O-rings</td><td>Grainger</td><td>1REC5</td><td>#010 Medium Hard Silicone O-Ring, 0.239&quot; I.D., 0.379&quot;O.D.</td></tr><tr><td>1/8&quot; Female luer to barb</td><td>Nordson biomedical</td><td>FTLL230-6005</td><td></td></tr><tr><td>1/8&quot; ID, 1/4&quot; OD autoclaveable PVC tubing</td><td>Thermo-Nalgene</td><td>63013-608</td><td>250&#039;</td></tr><tr><td>1/8&quot; male spinning luer to barb</td><td>Nordson biomedical</td><td>MLRL013-6005</td><td></td></tr><tr><td>1/8&quot; multiport barb</td><td>Nordson biomedical</td><td>4PLL230-6005</td><td>1/8&quot; multiport barb</td></tr><tr><td>1/8&quot; NPT to barb</td><td>Nordson biomedical</td><td>18230-6005</td><td>1/8&quot; 200 series barb</td></tr><tr><td>1/8&quot; panel mount luer</td><td>Nordson biomedical</td><td>Nordson MLRLB230-6005</td><td>1/8&quot;, PP</td></tr><tr><td>10 gallon fish tank</td><td>Walmart</td><td>802262</td><td>Can hold up to 8 bioreactors depending on layout</td></tr><tr><td>100-1000 ccm flow meter</td><td>Dwyer</td><td>RMA-13-SSV</td><td>For bottle reactors</td></tr><tr><td>2 ft fluorescent light bank</td><td>Agrobrite</td><td>FLT24 T5</td><td></td></tr><tr><td>200-2500 ccm flow meter</td><td>Dwyer</td><td>RMA-14-SSV</td><td>For air regulation upstream of humidifier</td></tr><tr><td>250 mL Pyrex bottles</td><td>Corning</td><td>16157-136</td><td>For gas mixing after humidifier</td></tr><tr><td>50-500 ccm flow meter</td><td>Dwyer</td><td>RMA-12-SSV</td><td>For hybridization tube reactors</td></tr><tr><td>5-50 ccm flow meter</td><td>Dwyer</td><td>RMA-151-SSV</td><td>For CO2 flow rate control</td></tr><tr><td>Air filters 0.2 &amp;micro;m</td><td>Whatman/ Fisher</td><td>09-745-1A</td><td>Polyvent, 28 mm, 0.2 &amp;micro;m, PTFE, 50 pack</td></tr><tr><td>Check valves</td><td>VWR</td><td>89094-714</td><td></td></tr><tr><td>Corning lids for pyrex bottles</td><td>VWR</td><td>89000-233</td><td>10 GL45 lids</td></tr><tr><td>Female luer endcap</td><td>Nordson biomedical</td><td>Nordson FTLLP-6005</td><td>Female stable PP</td></tr><tr><td>Hybridization tubes</td><td>Corning</td><td>32645-030</td><td>35x300 mm, pack of 2</td></tr><tr><td>Light timer</td><td>Walmart</td><td>556393626</td><td></td></tr><tr><td>Locknuts</td><td>Nordson biomedical</td><td>Nordson LNS-3</td><td>1/4&quot;, red nylon</td></tr><tr><td>Low profile magnetic stirrer</td><td>VWR</td><td>10153-690</td><td>Low profile magnetic stirrer</td></tr><tr><td>Male luer endcap</td><td>Nordson biomedical</td><td>Nordson LP4-6005</td><td>Male plug PP</td></tr><tr><td>Spinning luer lock ring</td><td>Nordson biomedical</td><td>Nordson FSLLR-6005</td><td></td></tr><tr><td>Stir bars - long</td><td>VWR</td><td>58949-040</td><td>38.1 mm, for bottle reactors</td></tr><tr><td>Stir bars - medium</td><td>VWR</td><td>58949-034</td><td>25 mm, for hyridization tubes</td></tr><tr><td>&lt;strong&gt;Supplies and reagents for culturing algae&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.2 &amp;micro;m filters</td><td>VWR</td><td>28145-491</td><td>13 mm, PTFE, for filtering spent media from daily culture sampling</td></tr><tr><td>1 mL syringes</td><td>Air-tite</td><td>89215-216</td><td>For filtering spent media from daily culture sampling</td></tr><tr><td>1.5 mL tubes</td><td>VWR</td><td>87003-294</td><td>Sterile (or equivalent)</td></tr><tr><td>10 mL Serological pipettes</td><td>Greiner Bio-One</td><td>82050-482</td><td>Sterile (or equivalent)</td></tr><tr><td>100 mm plates</td><td>VWR</td><td>25384-342</td><td>100x15 mm stackable petri dishes, sterile</td></tr><tr><td>15 mL tubes</td><td>Greiner Bio-One</td><td>82050-276</td><td>Sterile (or equivalent), polypropylene</td></tr><tr><td>2 mL Serological pipette tips</td><td>Greiner Bio-One</td><td>82051-584</td><td>Sterile (or equivalent)</td></tr><tr><td>2 mL tubes</td><td>VWR</td><td>87003-298</td><td>Sterile (or equivalent)</td></tr><tr><td>50 mL tubes</td><td>Greiner Bio-One</td><td>82050-348</td><td>Sterile (or equivalent), polypropylene</td></tr><tr><td>96 well microplate</td><td>Greiner Bio-One</td><td>89089-578</td><td>Polystyrene with lid, flat bottom</td></tr><tr><td>Inocculating loops</td><td>VWR</td><td>80094-478</td><td>Sterile (or equivalent)</td></tr><tr><td>Liquid carbon dioxide tank and regulator</td><td>Airgas</td><td>CD-50</td><td></td></tr><tr><td>&lt;strong&gt;Supplies and reagents for lipid extraction and neutral lipid assay&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>2 mL bead tubes</td><td>VWR</td><td>10158-556</td><td>Polypropylene tube w/ lid</td></tr><tr><td>96 well microplates</td><td>Greiner Bio-One</td><td>82050-774</td><td>Polypropylene, flat bottom</td></tr><tr><td>Bleach</td><td>Walmart</td><td>550646751</td><td>Only use regular bleach, not cleaning bleach</td></tr><tr><td>Chloroform</td><td>BDH</td><td>BDH1109-4LG</td><td></td></tr><tr><td>Dimethyl sulfoxide</td><td>BDH</td><td>BDH1115-1LP</td><td></td></tr><tr><td>Isopropyl alcohol</td><td>BDH</td><td>BDH1133-1LP</td><td></td></tr><tr><td>Methanol</td><td>BDH</td><td>BDH20864.400</td><td></td></tr><tr><td>Nile red</td><td>VWR</td><td>TCN0659-5G</td><td></td></tr><tr><td>Pasteur pipette tips</td><td>VWR</td><td>14673-010</td><td></td></tr><tr><td>Sodium chloride</td><td>BDH</td><td>BDH9286-500G</td><td></td></tr><tr><td>Vegetable oil</td><td>Walmart</td><td>9276383</td><td>Any vegetable oil should work as long as it is fresh</td></tr><tr><td>Zirconia/ silica beads (0.5 mm diameter)</td><td>Biospec products</td><td>11079105z</td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Analytical balance</td><td>Mettler-Toledo</td><td>XS205DU</td><td>Capable of at least 4 decimal accuracy</td></tr><tr><td>Bead homogenizer</td><td>Omni</td><td>19-040E</td><td></td></tr><tr><td>Benchtop micro centrifuge</td><td>Thermo</td><td>Heraeus Fresco 21 with 24x2</td><td>Including rotor capable of handling 1.5 and 2 mL tubes</td></tr><tr><td>Dry block heater</td><td>VWR</td><td>75838-282</td><td>Including dry block for a microplate</td></tr><tr><td>Freeze dryer</td><td>Labconco</td><td>7670520</td><td>2.5L freeze drying system</td></tr><tr><td>Large benchtop centrifuge</td><td>Thermo</td><td>Heraeus Megafuge 16R Tissue</td><td>Including rotors capable of handling 400 mL bottles, 50 mL tubes, and 15 mL tubes</td></tr><tr><td>Microplate reader</td><td>Molecular Devices</td><td>SpectraMax M2</td><td>Capable of reading absorbance and fluorescence</td></tr><tr><td>Vortex mixer</td><td>VWR</td><td>10153-838</td><td></td></tr></tbody></table>

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.

This procedure yields a time course of algal optical density data at OD 550 nm (Figure 4A). The optical density and dry weight concentration data can be correlated (Figure 4B). This is accomplished by first calculating the final dry weight algae concentration after the freeze-drying step. Next, the optical density of the culture serial dilution (performed on the last day of sampling) and the actual dry weight concentrations can b...

Watch this Scientific Journal Video about Cultivation of Green Microalgae in Bubble Column Photobioreactors and an Assay for Neutral Lipids at JoVE.com

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

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 ...

cultivation-green-microalgae-bubble-column-photobioreactors-an-assay

Universidad de Cartagena UDEC

Research

JoVE Journal

Environment

20.8K Views.  Auburn University. 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.

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

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 ...

A Simple and Rapid Protocol for Measuring Neutral Lipids in Algal Cells Using Fluorescence

Algae are considered excellent candidates for renewable fuel sources due to their natural lipid storage capabilities. Robust monitoring of algal fermentation processes and screening for new oil-rich strains requires a fast and reliable protocol for determination of intracellular lipid content. Current practices rely largely on gravimetric methods to determine oil content, techniques developed decades ago that are time consuming and require large sample volumes. In this paper, Nile Red, a fluorescent dye that has been used to identify the presence of lipid bodies in numerous types of organisms, is incorporated into a simple, fast, and reliable protocol for measuring the neutral lipid content of Auxenochlorella protothecoides, a green alga. The method uses ethanol, a relatively mild solvent, to permeabilize the cell membrane before staining and a 96 well micro-plate to increase sample capacity during fluorescence intensity measurements. It has been designed with the specific application of monitoring bioprocess performance. Previously dried samples or live samples from a growing culture can be used in the assay.

A simple protocol to determine the neutral lipid content of algal cells using a Nile Red staining procedure is described. This time-saving technique offers an alternative to traditional gravimetric-based lipid quantification protocols. It has been designed for the specific application of monitoring bioprocess performance.

Algae are considered excellent candidates for renewable fuel sources due to their natural lipid storage capabilities. Robust monitoring of algal ...

Chemistry

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 ...

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 ...

Coupling Carbon Capture from a Power Plant with Semi-automated Open Raceway Ponds for Microalgae Cultivation

In the United States, 35% of the total carbon dioxide (CO2) emissions come from the electrical power industry, of which 30% represent natural gas electricity generation. Microalgae can biofix CO2 10 to 15 times faster than plants and convert algal biomass to products of interest, such as biofuels. Thus, this study presents a protocol that demonstrates the potential synergies of microalgae cultivation with a natural gas power plant situated in the southwestern United States in a hot semi-arid climate. State-of-the-art technologies are used to enhance carbon capture and utilization via the green algal species Chlorella sorokiniana, which can be further processed into biofuel. We describe a protocol involving a semi-automated open raceway pond and discuss the results of its performance when it was tested at the Tucson Electric Power plant, in Tucson, Arizona. Flue gas was used as the main carbon source to control pH, and Chlorella sorokiniana was cultivated. An optimized medium was used to grow the algae. The amount of CO2 added to the system as a function of time was closely monitored. Additionally, other physicochemical factors affecting algal growth rate, biomass productivity, and carbon fixation were monitored, including optical density, dissolved oxygen (DO), electroconductivity (EC), and air and pond temperatures. The results indicate that a microalgae yield of up to 0.385 g/L ash-free dry weight is attainable, with a lipid content of 24%. Leveraging synergistic opportunities between CO2 emitters and algal farmers can provide the resources required to increase carbon capture while supporting the sustainable production of algal biofuels and bioproducts.

A protocol is described to utilize the carbon dioxide in natural gas power plant flue gas to cultivate microalgae in open raceway ponds. Flue gas injection is controlled with a pH sensor, and microalgae growth is monitored with real time measurements of optical density.

In the United States, 35% of the total carbon dioxide (CO2) emissions come from the electrical power industry, of which 30% represent natural gas ...

Microalgal Cultivation for Biomethane Obtention from Swine Waste Biogas

Biogas Purification through the use of a Microalgae-Bacterial System in Semi-Industrial High Rate Algal Ponds

In recent years, a number of technologies have emerged to purify biogas into biomethane. This purification entails a reduction in the concentration of polluting gases such as carbon dioxide and hydrogen sulfide to increase the content of methane. In this study, we used a microalgal cultivation technology to treat and purify biogas produced from organic waste from the swine industry to obtain ready-to-use biomethane. For cultivation and purification, two 22.2 m3 open-pond photobioreactors coupled with an absorption-desorption column system were set up in San Juan de los Lagos, Mexico. Several recirculation liquid/biogas ratios (L/G) were tested to obtain the highest removal efficiencies; other parameters, such as pH, dissolved oxygen (DO), temperature, and biomass growth, were measured. The most efficient L/Gs were 1.6 and 2.5, resulting in a treated biogas effluent with a composition of 6.8%vol and 6.6%vol in CO2, respectively, and removal efficiencies for H2S up to 98.9%, as well as maintaining O2 contamination values of less than 2%vol. We found that pH greatly determines CO2 removal, more so than L/G, during cultivation because of its participation in the photosynthetic process of microalgae and its ability to vary pH when solubilized due to its acidic nature. DO, and temperature oscillated as expected from the light-dark natural cycles of photosynthesis and the time of day, respectively. Biomass growth varied with CO2 and nutrient feeding as well as reactor harvesting; however, the trend remained primed for growth.

Author Spotlight: Scaling Microalgal Biotechnology for Enhanced Biomethane Production 

Air pollution impacts the quality of life of all organisms. Here, we describe the use of microalgae biotechnology for the treatment of biogas (simultaneous removal of carbon dioxide and hydrogen sulfide) and the production of biomethane through semi-industrial open high-rate algal ponds and subsequent analysis of treatment efficiency, pH, dissolved oxygen, and microalgae growth.

In recent years, a number of technologies have emerged to purify biogas into biomethane. This purification entails a reduction in the concentration of ...

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 ...

Bioengineering

Enhancing Chlorophyll Production in Microalgae via Growth Factor Optimization

Evaluation of the Effect of Growth Factors on Chlorophylls a and b Production from Microalgae

Microalgae contain two main groups of pigments: chlorophylls and carotenoids. Chlorophyll is a green pigment that absorbs light energy and transforms it into chemical energy to facilitate the synthesis of organic compounds. This pigment serves as a valuable primary source for biotechnological input products in the food, pharmaceutical, and cosmetic industries due to its high antioxidant properties and coloring capabilities. The objective of this research was to evaluate the effect of growth factors (CO2 concentration, light color, and light intensity) through a Taguchi L4 experimental design on cell growth and the cellular content of chlorophyll a and b in Chlorella sorokiniana, followed by validation of the method using Haematococcus pluvialis microalgae as an additional study model. Cell growth was quantified using the optical density spectrophotometric technique at a wavelength of 550 nm. For the quantification of chlorophylls, a cell extract was obtained using a 90% pure acetone solution, and subsequently, the concentrations of chlorophylls a and b were quantified using spectrophotometric techniques at wavelengths of 647 nm and 664 nm, according to the method described by Jeffrey and Humphrey. The experimental results indicated that controlling conditions of low CO2 addition, purple light, and low light intensity increases both cell growth and the concentration of chlorophylls a and b within the cells. The implementation of this chlorophyll quantification method allows for quick, simple, and precise determination of chlorophyll content, as the wavelengths used are at the absorbance peaks of both types of chlorophylls, making this technique easily reproducible for any microalgae under study.

Author Spotlight: Optimizing Growth Factors for Production of Biotechnologically Relevant Secondary Metabolites

The present study highlights the advantages of employing the method developed by Jeffrey and Humphrey for extracting and quantifying fat-soluble pigments from microalgae. This method serves as a valuable tool for assessing the influence of growth factors on chlorophyll production and cellular content in these organisms.

Microalgae contain two main groups of pigments: chlorophylls and carotenoids. Chlorophyll is a green pigment that absorbs light energy and transforms it ...

Biochemistry

High-Throughput Metabolic Profiling for Model Refinements of Microalgae

Metabolic models are reconstructed based on an organism&#39;s available genome annotation and provide predictive tools to study metabolic processes at a systems-level. Genome-scale metabolic models may include gaps as well as reactions that are unverified experimentally. Reconstructed models of newly isolated microalgal species will result in weaknesses due to these gaps, as there is usually sparse biochemical evidence available for the metabolism of such isolates. The phenotype microarray (PM) technology is an effective, high-throughput method that functionally determines cellular metabolic activities&#160;in response to a wide array of entry metabolites. Combining the high throughput phenotypic assays with metabolic modeling can allow existing metabolic network models to be rapidly reconstructed or optimized by providing biochemical evidence to support and expand genomic evidence. This work will show the use of PM assays for the study of microalgae by using the green microalgal model species Chlamydomonas reinhardtii as an example. Experimental evidence for over 254 reactions obtained by PM was used in this study to expand and refine a genome-scale C. reinhardtii metabolic network model, iRC1080, by approximately 25 percent. The protocol created here can be used as a basis for functionally profiling the metabolism of other microalgae, including known microalgae mutants and new isolates.

This protocol demonstrates the use of a phenotype microarray (PM) technology platform to define metabolic requirements of Chlamydomonas reinhardtii, a green microalga, and refine an existing metabolic network model.

Metabolic models are reconstructed based on an organism&#39;s available genome annotation and provide predictive tools to study metabolic processes at a ...