The authors gratefully acknowledge the partial funding from the TecNM under the Call for Scientific Research, Technological Development, and Innovation (16898.23-P) for the Institutos Tecnologicos Federales. They also appreciate the support from the Instituto de Ciencia, Tecnolog&#237;a e Innovaci&#243;n del Estado de Michoac&#225;n de Ocampo (FCCHTI23_ME-4.1.-0001).

1. Culture media preparation and inoculum preparation
<ol>
	<li>Prepare 1 L of 3N-BBM+ V(CCAP) growth medium (Macronutrients: NaNO3 [0.75 g L-1], CaCl2 [0.019 g L-1], MgSO4 [0.019 g L-1], K2HPO4 [0.057 g L-1], NaCl [0.025 g L-1], KH2PO4 [0.175 g L-1]; micronutrients: Na2 EDTA [0.0186 g L-1], FeCl3 [0.0024 g L-1], MnCl2 [0.001 g L-1], ZnCl2 [1.25 x 10-4 g L-1], CoCl2 [5 x 10-5 g L-1], Na2MoO4 [9.98 x 10-5 g L-1])8 and sterilize it for 15 min at 121 &#176;C and 1.5 psi.</li>
	<li>Inoculate a flask with 1 L of growth medium at an approximate concentration of 3 x 106 cells mL-1 for the microalga C. sorokiniana; or add 5% to 10% (v/v) of inoculum to the growth medium. 
	NOTE: To maintain sterile conditions, it is recommended to use a laminar flow hood, gloves, lab coat, and face mask.</li>
	<li>Incubate the inoculated flask under red light sources with an approximate intensity of 3,600 to 4,200 lux, at a temperature between 22-26 &#176;C, with a photoperiod of 12 h light and 12 h darkness for C. sorokiniana and photoperiod of 16 h light and 8 h darkness for H. pluvialis, and manually agitate every 24 h, approximately for 7 days for both strains. 
	NOTE: Maintain under growth conditions for at least 7-10 days (until obtaining an intense green color) for C. sorokiniana and 10-12 days for H. pluvialis. To obtain the microalgae concentrate that will serve as the inoculum in the kinetics, it is necessary to remove the supernatant medium from the produced algae.</li>
	<li>In a sterile environment, fill sterile 15 mL conical centrifugation tubes with approximately 10 mL of inoculated medium and centrifuge at 3,000 x g for 20 min under 25-30 &#176;C.</li>
	<li>Under sterile conditions, remove the supernatant using a micropipette or by pouring and concentrating the biomass in a sterile 50 mL conical centrifugation tube.</li>
	<li>Repeat this process as necessary until the required volume of inoculum for the kinetics is obtained.</li>
	<li>Measure the optical density (OD550&#160;nm) of the inoculum to determine the cell concentration of the microalgae and store it at a temperature of 3-4 &#176;C until ready for use. 
	NOTE: Direct cellular counting was initially performed to correlate cell concentration with absorbance. Once counted, the inoculum can be stored under refrigeration for no more than 4 days.</li>
</ol>
2. Study of the growth kinetics
<ol>
	<li>Prepare the required volume of 3N-BBM+ V(CCAP) suitable for each experiment (8 L for photobioreactor level tests for C. sorokiniana and 0.5 L for the flask-level tests for H. pluvialis) according to standard laboratory procedures, ensuring proper sterilization.</li>
	<li>For both photobioreactor and flasks, configure the light source to the desired color spectrum (Blue at 460 nm, purple, mix of 460 nm and 630 nm for C. sorokiniana and red at 630 nm for H. pluvialis); cover both systems to avoid external light interference. 
	NOTE: Due to the variation in wavelength emitted from each light-emitting diode, spectroradiometry was performed using a spectroradiometer on the light sources to determine the wavelength and emission power of each diode9. Purple light was achieved by combining red and blue LEDs, resulting in wavelengths of 460 nm with an emission power of 1.65 W nm-1&#160;for the blue LEDs and 630 nm with an emission power of 0.6 W nm-1&#160;for the red LEDs. These measurements were used for the tests conducted solely under blue and red light.</li>
	<li>Program the timer to alternate between 12 h light and 12 h dark cycles for C. sorokiniana and 16 h light and 8 h dark cycles for H. pluvialis.</li>
	<li>Connect the CO2 aeration system to the photobioreactor culture vessel to provide carbon dioxide supply in intervals of 12 h for C. sorokiniana.</li>
	<li>Inoculate the culture vessel with a sufficient volume of pre-inoculum to achieve a starting cell density of 3 x 105 cells mL-1 or its equivalent in optical density (OD550&#160;nm) = 0.180. 
	NOTE: Ensure aseptic technique during inoculation to prevent contamination.</li>
	<li>Sample the culture at regular intervals of 12 h for C. sorokiniana and 8 h for H. pluvialis to monitor cell growth and physiological parameters. Collect samples using sterile techniques to maintain the integrity of the culture.</li>
</ol>
3. Quantification of chlorophylls
NOTE: Data for cellular quantification is provided in Supplementary File 1.
<ol>
	<li>Place 40 mL of the sample into plastic conical centrifugation tubes.</li>
	<li>Centrifuge at 3,000 x g for 20 min at 15 &#176;C and discard the supernatant by decantation or using a Pasteur pipette.</li>
	<li>Transfer the cell pellet into a glass tube covered with aluminum foil and a screw cap to prevent oxidation.</li>
	<li>Resuspend the cell pellet with 3 mL of 90% pure acetone solution using a vortex.</li>
	<li>Sonicate the suspended sample in an ice bath for two cycles of 5 min each and let it rest at 4 &#176;C for 16 h10.</li>
	<li>After 16 h, sonicate again under the same conditions mentioned in the previous step (step 3.5) and centrifuge again under the previously described conditions (step 3.2).</li>
	<li>Separate the pigment extract using a Pasteur pipette and transfer it to another clean tube protected from light.</li>
	<li>Place the pigment extract in a quartz cell to read in a spectrophotometer at 630 nm, 647 nm, and 664 nm wavelengths. 
	NOTE: Calibrate the spectrophotometer beforehand with 90% acetone.</li>
	<li>To calculate chlorophyll concentrations, use the following equations11: 
	Chlorophyll a = 11.93 A664 - 1.93 A647 
	Chlorophyll b = 20.36 A647 - 5.50 A664 
	NOTE: Results are expressed in &#181;g mL-1 of extract, which are multiplied by the extract volume and divided by the number of mL of sample to obtain chlorophyll concentrations in &#181;g mL-1 of culture.</li>
	<li>To obtain values in &#181;g chl cell-1, use the following formula: 
	&#956;g chl cell-1 = &#181;g chl in 1 mL of culture / [cells mL-1] cells concentration in the culture.</li>
</ol>

Enhancing Chlorophyll Production in Microalgae via Growth Factor Optimization

<ol>
	<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> Microb Cell Fact. 17 (1), 36 (2018).</li><li>Otero-Paternina, A. S. M., Cruz-Casallas, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+the+hydrocarbon+phenanthrene+on+Chlorella+vulgaris+(Chlorellaceae)+growth.">Effect of the hydrocarbon phenanthrene on Chlorella vulgaris (Chlorellaceae) growth.</a> Acta Biol Col. 18 (1), 87-98 (2013).</li><li>Pancha, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Salinity-induced+oxidative+stress+enhanced+the+biofuel+production+potential+of+microalgae+Scenedesmus+sp.+CCNM+1077.">Salinity-induced oxidative stress enhanced the biofuel production potential of microalgae Scenedesmus sp. CCNM 1077.</a> Bioresour Techno. 189, 341-348 (2015).</li><li>Ortiz, M. L., Cort&#233;s, C. E., S&#225;nchez, J., Padilla, J., Otero, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluating+microalgae+Chlorella+sorokiniana+growth+in+different+culture+mediums+in+autotrophic+and+mixotrophic+conditions.">Evaluating microalgae Chlorella sorokiniana growth in different culture mediums in autotrophic and mixotrophic conditions.</a> Orinoquia. 16 (1), 11-20 (2012).</li><li>Camacho Kurmen, J. E., Gonz&#225;lez, G., Klotz, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astaxanthin+production+in+Haematococcus+pluvialis+under+different+stress+conditions.">Astaxanthin production in Haematococcus pluvialis under different stress conditions.</a> Nova. 11 (19), 94-104 (2013).</li><li>. Production of chlorophyll and astaxanthin from Haematococcus pluvialis under nitrogen deficiency stress in a 5-liter Biostat Aplus bioreactor Available from: <a target="_blank" href="https://repositorio.unicolmayor.edu.co/handle/unicolmayor/2735">https://repositorio.unicolmayor.edu.co/handle/unicolmayor/2735</a> (2019)</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=Phycological+studies+IV.+Some+soil+algae+from+Enchanted+Rock+and+related+algal+species.">Phycological studies IV. Some soil algae from Enchanted Rock and related algal species.</a> University of Texas. 6318, 95 (1963).</li><li>Cerezo, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=LED+characterization+through+the+use+and+programming+of+a+spectrometer.">LED characterization through the use and programming of a spectrometer.</a> Universidad Polit&#233;cnica de. 22323, (2013).</li><li>Couso, I., Vila, M., Vigara, J., Cordero, B. F., Vargas, M. &. #. 1. 9. 3. ;. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synthesis+of+carotenoids+and+regulation+of+the+carotenoid+biosynthesis+pathway+in+response+to+high+light+stress+in+the+unicellular+microalga+Chlamydomonas+reinhardtii.">Synthesis of carotenoids and regulation of the carotenoid biosynthesis pathway in response to high light stress in the unicellular microalga Chlamydomonas reinhardtii.</a> Eur J Phycol. 47 (3), 223-232 (2012).</li><li>Vera-L&#243;pez, F., Mart&#237;nez, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pigments+in+microalgae:+Functions,+applications,+and+overproduction+techniques.">Pigments in microalgae: Functions, applications, and overproduction techniques.</a> BioTechnology. 25 (5), 35-51 (2021).</li></ol>

The authors have nothing to disclose.

The comparative study between H. pluvialis and C. sorokiniana revealed significant differences in chlorophyll production dynamics. While H. pluvialis exhibited a decrease in chlorophyll concentration throughout the experiment, C. sorokiniana showed a steady increase. Additionally, there was initially a lower proportion of chlorophyll a in both species, but this ratio reversed in particular growth conditions, which may give indications of an induction of the production of said ...

In recent years, the growing environmental problems caused by anthropogenic activities and their adverse effects on the health and balance of ecosystems have driven the search for more efficient and environmentally friendly production systems. This has accelerated processes in industries and fostered the implementation of bioremediation treatments and the development of bio compounds to mitigate these harmful effects1.
This context has led to a significant growth in the study of microalgae, driven by the need to find innovative solutions to current environmental and economic challenges. Microalgae thrive in aquatic environments, using sunlight and carbon dioxide as sources of energy and carbon, respectively. This characteristic makes them a sustainable and promising alternative for producing various valuable products. Research in this field has focused on understanding the physiology and metabolism of these cells, as well as developing efficient technologies for their cultivation and processing2.
There is a clear need to develop accessible and reliable tools for studying microalgae to accelerate research processes and deepen the understanding of their physiology, metabolism, and potential applications. These tools should enable rapid analysis of the effect of environmental and production factors on key parameters, such as chlorophyll concentration, which is a fundamental indicator of the health and development of these organisms. A decrease in chlorophyll content may indicate environmental stress, nutritional deficiencies, or diseases.
These green pigments play a crucial role in photosynthesis, capturing sunlight energy to convert carbon dioxide and water into glucose and oxygen and constitute between 0.5% and 5% of the biomass of microalgae3. Beyond their essential role in sustaining life processes, chlorophylls have found applications in various industries. Chlorophyll extracts are utilized in food and beverage processing as natural colorants, imparting vibrant green hues to products while also providing antioxidant properties. Moreover, chlorophyll-based supplements are gaining popularity in the health and wellness sector due to their purported detoxifying and anti-inflammatory effects. By harnessing the multifaceted properties of chlorophyll, industries can develop innovative products that contribute to both visual appeal and consumer well-being1.
In this regard, one microalgae of great biotechnological relevance is C. sorokiniana. This microorganism stands out for its rapid growth rate, making it highly efficient in biomass production. Additionally, C. sorokiniana exhibits a diverse content of highly nutritional compounds, including proteins, lipids, and vitamins, which renders it valuable for various applications in food, feed, and biofuel production2. Moreover, this microalgae species has been found to produce extracellular enzymes with diverse functions, opening up possibilities for biotechnological applications such as wastewater treatment, bioremediation, and pharmaceuticals4. In addition to its rapid growth and versatile applications, C. sorokiniana also demonstrates significant potential for chlorophyll production. As a photosynthetic microorganism, C. sorokiniana possesses the machinery necessary for the synthesis of chlorophyll, which constitutes between 0.5% and 5% of the biomass of microalgae3. This capability makes C. sorokiniana an attractive candidate for commercial-scale chlorophyll production and promises to be a sustainable solution to urgent environmental and nutritional challenges5.
On the other hand, another microalgae species of significant interest is H. pluvialis. This microalga is renowned for its production of astaxanthin, a potent antioxidant pigment with numerous industrial applications. Astaxanthin serves as a protective mechanism for H. pluvialis photosystem, shielding it from oxidative stress induced by environmental factors. This pigment is highly sought after in cosmetics, nutraceuticals, and aquaculture industries, owing to its antioxidant properties and potential health benefits. With its abundant ability to produce astaxanthin, H. pluvialis presents a promising avenue for developing innovative products catering to various industrial and consumer needs6.
Recent studies have also highlighted H. pluvialis1 potential for chlorophyll production, further enhancing its industrial significance. For instance, a study investigated the chlorophyll content of H. pluvialis under varying growth conditions and found that under optimal cultivation parameters, H. pluvialis exhibited remarkable chlorophyll production rates, surpassing those of other microalgae species7. This finding underscores the potential of H. pluvialis as a sustainable source of chlorophyll, offering novel opportunities for its utilization in various industrial sectors.

<table><tbody><tr><td>C&lt;sub&gt;3&lt;/sub&gt;H&lt;sub&gt;6&lt;/sub&gt;O</td><td>Meyer</td><td>67-64-1</td><td>Acetone 90%</td></tr><tr><td>15 mL tube</td><td>Biologix</td><td>10-9502</td><td>Test tube</td></tr><tr><td>2510-DTH</td><td>Branson</td><td>D-73595</td><td>Sonicator</td></tr><tr><td>5 mL screw cap test tube</td><td>Kimax</td><td>45066-13100</td><td>Test tube</td></tr><tr><td>50 mL centrifuge tube</td><td>Biologix</td><td>10-9151</td><td>Test tube</td></tr><tr><td>Aluminum foil</td><td>Reynolds</td><td>611 standard, 12&quot; x 1000 feet</td><td>Test tube cover&amp;nbsp;</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Meyer</td><td>0925-250</td><td>Calcium Chloride</td></tr><tr><td>Centrifuge&amp;nbsp;</td><td>Dynamica</td><td>14 R</td><td>Centrifuge Refrigerated</td></tr><tr><td>CoCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Merck</td><td>1057-100</td><td>Cobalt dichloride</td></tr><tr><td>FeCl&lt;sub&gt;3&lt;/sub&gt;</td><td>Merck</td><td>157740</td><td>Iron(III) Chloride</td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Meyer</td><td>2051-250</td><td>Dipotassium Phosphate</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Meyer</td><td>2055-250</td><td>Monopotassium Phosphate</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Meyer</td><td>1605-250</td><td>Magnesium Sulphate</td></tr><tr><td>Micropipette</td><td>LabNet</td><td>Model Beta-Pette</td><td>Micropipette</td></tr><tr><td>MnCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Merck</td><td>429449</td><td>Manganese(II) Chloride&amp;nbsp;</td></tr><tr><td>Na2 EDTA&amp;nbsp;</td><td>Merck</td><td>200-449-4</td><td>Edatamil, Edetato Disodium Salt Dihydrate</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;MoO&lt;sub&gt;4&lt;/sub&gt;</td><td>Merck</td><td>243655</td><td>Sodium Molybdate</td></tr><tr><td>NaCl</td><td>Meyer</td><td>2365-500</td><td>Sodium Chloride</td></tr><tr><td>NaNO&lt;sub&gt;3&lt;/sub&gt;</td><td>Meyer</td><td>2465-250</td><td>Sodium Nitrate</td></tr><tr><td>RGB LED stripe</td><td>Steren</td><td>GAD-LED2</td><td>Light source</td></tr><tr><td>Spectrophotometer</td><td>PerkinElmer</td><td>Model Lambda35</td><td>Spectrophotometer</td></tr><tr><td>spectroradiometer</td><td>Gigahertz-Optik</td><td>model BTS256</td><td></td></tr><tr><td>Vortex</td><td>Scientific Industries</td><td>Vortex-Genie&amp;reg; 2</td><td>Vortex</td></tr><tr><td>ZnCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Merck</td><td>208086</td><td>Zinc Chloride</td></tr></tbody></table>

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.

To observe the efficiency of the technique detecting variations in chlorophyll cellular concentration and evaluate the effect of growth factors in C. sorokiniana, a Taguchi L4 experimental design was established, evaluating CO2 volume addition, light color, and light intensity. Each factor was assessed at low and high levels, as shown in Table 1, under the conditions defined by the experimental design in Table 2.
Once the experim...

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.

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 ...

evaluation-effect-growth-factors-on-chlorophylls-b-production-from

Research

JoVE Journal

Biochemistry

2.1K Views.  National Institute of Technology of Mexico. Ons project evalueert hoe groeifactorcontrole de productie van biotechnologisch relevante secundaire metabolieten zoals chlorofyl uit microalgen zoals Chlorella sorokiniana en Haematococcus pluvialis op verschillende schalen beïnvloedt. We hebben ons gericht op het opzetten van een analytische methode om chlorofyl in microalgen te kwantificeren. Enkele opkomende technologieën om hierin vooruitgang te boeken, zijn het modelleren en optimaliseren van processe...

Video: Auteur in de schijnwerpers: Optimalisatie van groeifactoren voor de productie van biotechnologisch relevante secundaire metabolieten

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 ...

Environment

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 Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated (WATER-PAM) Fluorometry

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be subsampled throughout the experiment1&#8211;7. Subsampling of large volumes can be problematic for several reasons: 1) it causes variation in the total volume and the surface area:volume ratio of the culture during the experiment; 2) pseudo-replication (i.e., replicate samples from the same treatment flask8) is often employed rather than true replicates (i.e., sampling from replicate treatments); 3) the duration of the experiment is limited by the total volume; and 4) axenic cultures or the usual bacterial microbiota are difficult to maintain during long-term experiments as contamination commonly occurs during subsampling.
The use of microtiter plates enables 1 ml culture volumes to be used for each replicate, with up to 48 separate treatments within a 12.65 x 8.5 x 2.2 cm plate, thereby decreasing the experimental volume and allowing for extensive replication without subsampling any treatment. Additionally, this technique can be modified to fit a variety of experimental formats including: bacterial-algal co-cultures, algal physiology tests, and toxin screening9&#8211;11. Individual wells with an alga, bacterium and/or co-cultures can be sampled for numerous laboratory procedures including, but not limited to: WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry, microscopy, bacterial colony forming unit (cfu) counts and flow cytometry. The combination of the microtiter plate format and WATER-PAM fluorometry allows for multiple rapid measurements of photochemical yield and other photochemical parameters with low variability between samples, high reproducibility and avoids the many pitfalls of subsampling a carboy or conical flask over the course of an experiment.

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated WATER-PAM Fluorometry

The goal of this procedure is to demonstrate the reproducibility and adaptability of using a microtiter plate format for microalgal screening. This rapid screen combines WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry to measure photosynthetic yield as an indicator of Photosystem II (PSII) health with small volume bacterial-algal co-cultures.

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be ...

Comparison of Scale in a Photosynthetic Reactor System for Algal Remediation of Wastewater

An experimental methodology is presented to compare the performance of two different sized reactors designed for wastewater treatment. In this study, ammonia removal, nitrogen removal and algal growth are compared over an 8-week period in paired sets of small (100 L) and large (1,000 L) reactors designed for algal remediation of landfill wastewater. Contents of the small and large scale reactors were mixed before the beginning of each weekly testing interval to maintain equivalent initial conditions across the two scales. System characteristics, including surface area to volume ratio, retention time, biomass density, and wastewater feed concentrations, can be adjusted to better equalize conditions occurring at both scales. During the short 8-week representative time period, starting ammonia and total nitrogen concentrations ranged from 3.1-14 mg NH3-N/L, and 8.1-20.1 mg N/L, respectively. The performance of the treatment system was evaluated based on its ability to remove ammonia and total nitrogen and to produce algal biomass. Mean &#177; standard deviation of ammonia removal, total nitrogen removal and biomass growth rates were 0.95&#177;0.3 mg NH3-N/L/day, 0.89&#177;0.3 mg N/L/day, and 0.02&#177;0.03 g biomass/L/day, respectively. All vessels showed a positive relationship between the initial ammonia concentration and ammonia removal rate (R2=0.76). Comparison of process efficiencies and production values measured in reactors of different scale may be useful in determining if lab-scale experimental data is appropriate for prediction of commercial-scale production values.

An experimental methodology is presented to compare the performance of small (100 L) and large (1,000 L) scale reactors designed for algae remediation of landfill wastewater. System characteristics, including surface area to volume ratio, retention time, biomass density, and wastewater feed concentrations, can be adjusted based on application.

An experimental methodology is presented to compare the performance of two different sized reactors designed for wastewater treatment. In this study, ...

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

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

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

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

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

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

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

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

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 ...

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

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 ...

Biology

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

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

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

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

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

Samenvatting

Meer video's verkennen

Hoofdstukken in deze video