A part of this work was financially supported by the Institute for the Promotion of Innovation by Science and Technology&mdash;Strategic Basic Research (IWT-SBO) project Sunlight and Biosolar cells. Erik Bolder and BackKim Nguyen are acknowledged for their contribution to the optimization of the bead beating procedure.

1. Sample Preparation
There are two alternate protocols for sample preparation included as steps 1.1 and 1.2. Both methods are equally suitable and give similar results, but if a limited amount of algae culture volume is available, method 1.1 is recommended.
NOTE: For either protocol, prepare two additional bead beater tubes according to the entire protocol but without adding algae to them to be used as a blank. In this way, peaks in the GC chromatogram resulting from extraction of components from materials used can be identified and quantified.
1.1. Sample Preparation Protocol Option 1: Recommended When a Limited Amount of Algae Culture is Available
<ol>
	<li>Determine the algae dry weight concentration (g/L) in the culture broth, for example as described by Breuer et al.1.</li>
	<li>Transfer a volume of culture broth that contains 5-10 mg algae dry weight to a glass centrifuge tube. Calculate the exact amount of biomass transferred using the biomass concentration determined in step 1.1.1.</li>
	<li>Centrifuge 5 min at 1,200 x g.</li>
	<li>Discard part of the supernatant, leave approximately 0.25 ml in the tube.</li>
	<li>Re-suspend the algae in the remaining supernatant by gentle pipetting the pellet up and down and transfer the complete cell pellet to a bead beater tube using a 200 &mu;l pipette.</li>
	<li>Rinse the centrifuge tube and glass pipette with &plusmn;0.15 ml milliQ water and transfer the liquid to the same bead beater tube.</li>
	<li>Centrifuge bead beater tubes for one minute at maximum rpm to make sure no air bubbles remain in the bottom of the tubes. It is possible to store closed bead beater tubes at -80 &deg;C.</li>
	<li>Lyophilize the bead beater tubes containing the sample. It is possible to store the closed bead beater tubes at -80 &deg;C.</li>
</ol>
1.2. Sample Preparation Protocol Option 2
<ol>
	<li>Centrifuge an undetermined volume of algae culture broth and discard supernatant. It is not necessary to determine the biomass concentration or to measure the volume used.</li>
	<li>Measure or calculate the osmolarity of the culture medium using the concentration of the main salts present in the culture medium.</li>
	<li>Wash the cell pellet by resuspending the cell pellet in the same volume, as used in step 1.2.1, of an ammonium formate solution, which approximates the osmolarity of the culture medium. An equiosmolar ammonium formate solution prevents cell lyses during washing of the cells.</li>
</ol>
NOTE: Washing the cells is necessary to remove salts present in the culture medium. This is especially important for fatty acid analysis in marine microalgae because of the high salt concentrations in their culture medium. If salts would still be present, this would cause overestimation of the amount of cell dry weight which will be determined later in this protocol. Ammonium formate is used to wash cells because this solution will completely evaporate during lyophilization and not leave any residue.
<ol start="4">
	<li>Centrifuge algae suspension and discard supernatant.</li>
	<li>Lyophilize the cell pellet.</li>
	<li>Weigh 5-10 mg lyophilized microalgae powder into a bead beater tube. Record the exact weight.</li>
</ol>
2. Cell Disruption and Lipid Extraction
NOTE: This section describes an extensive cell disruption procedure. Possibly, some of the cell disruption steps are redundant and less extensive cell disruption might yield the same results for some microalgae species or for biomass derived from certain cultivation conditions. This would need validation for each specific situation however. Therefore the proposed extensive disruption protocol is recommended as a universal method suitable for all microalgal material.
<ol>
	<li>Weigh an exact amount of tripentadecanoin (triacylglycerol containing three C15:0 fatty acids) and add it to an exactly known amount of 4:5 (v/v) chloroform:methanol. In this way the concentration of tripentadecanoin is exactly known. Aim for a concentration of 50 mg/L. Tripentadecanoin serves as the internal standard for fatty acid quantification.</li>
	<li>Add 1 ml chloroform:methanol 4:5 (v/v) containing tripentadecanoin to each beating tube (specified in the reagents and equipment table). Check that there are no beads remaining inside the cap of the beadbeater tube, this will result in leaking of the tubes. Use a positive displacement pipette for accurate addition of solvents.</li>
	<li>Bead beat the bead beater tubes 8x at 2,500 rpm for 60 sec, each time with a 120 sec internal between each beating.</li>
	<li>Transfer the solution from the bead beater tubes to a clean heat-resistant 15 ml glass centrifuge tube with Teflon insert screw-caps. Be sure to transfer all the beads from the bead beater tube as well.</li>
	<li>To wash the bead beater tube, add 1 ml chloroform:methanol 4:5 (v/v) containing tripentadecanoin into the bead beater tube, close cap and mix, and transfer this solution to the same glass tube from step 2.4. Wash the tube three times (in total 4 ml of chloroform:methanol 4:5 (v/v) containing tripentadecanoin is used per sample - 1 ml added in step 2.2 and 3 ml for 3 washing steps).</li>
	<li>Vortex the glass tubes for 5 sec and sonicate in a sonication bath for 10 min.</li>
	<li>Add 2.5 ml of MilliQ water, containing 50 mM 2-amino-2-hydroxymethyl-propane-1,3-diol (Tris) and 1 M NaCl, which is set to pH 7 using a HCl solution. This will cause a phase separation between chloroform and methanol:water. 1 M NaCl is used to enhance the equilibrium of lipids towards the chloroform phase.</li>
	<li>Vortex for 5 sec and then sonicate for 10 min.</li>
	<li>Centrifuge for 5 min at 1,200 x g.</li>
	<li>Transfer the entire chloroform phase (bottom phase) to a clean glass tube using a glass Pasteur pipette. Make sure not to transfer any interphase or top phase.</li>
	<li>Re-extract the sample by adding 1 ml chloroform to old tube (containing the methanol:water solution).</li>
	<li>Vortex for 5 sec and then sonicate for 10 min.</li>
	<li>Centrifuge for 5 min at 1,200 x g.</li>
	<li>Collect the chloroform phase (bottom phase) using a glass Pasteur pipette and pool with the first chloroform fraction from step 2.10.</li>
	<li>If difficulties are experienced in collecting the entire chloroform fraction in the previous step, repeat steps 2.11-2.14. Otherwise proceed with step 2.16.</li>
	<li>Evaporate the chloroform from the tube in a nitrogen gas stream. After this step samples can be stored at -20 &deg;C under a nitrogen gas atmosphere.</li>
</ol>
3. Transesterification to Fatty Acid Methyl Esters
<ol>
	<li>Add 3 ml methanol containing 5% (v/v) sulfuric acid to the tube containing the dried extracted lipids (tube originally containing the chloroform fraction) and close the tube tightly.</li>
	<li>Vortex for 5 sec.</li>
	<li>Incubate samples for 3 hr at 70 &deg;C in a block heater or water bath. Periodically (every &plusmn;30 min) ensure that the samples are not boiling (caused by an improperly closed lid) and vortex tubes. During this reaction fatty acids are methylated to their fatty acid methyl esters (FAMEs).</li>
	<li>Cool samples to room temperature and add 3 ml MilliQ water and 3 ml n-hexane.</li>
	<li>Vortex for 5 sec and mix for 15 min with a test tube rotator.</li>
	<li>Centrifuge for 5 min at 1,200 x g.</li>
	<li>Collect 2 ml of the hexane (top) phase and put in fresh glass tube using a glass Pasteur pipette.</li>
	<li>Add 2 ml MilliQ water to the collected hexane phase to wash it.</li>
	<li>Vortex for 5 sec and centrifuge for 5 min at 1,200 x g. After this step it is possible to store the samples at -20 &deg;C under nitrogen gas atmosphere (phase separation not necessary).</li>
</ol>
4. Quantification of FAMEs Using Gas Chromatography
<ol>
	<li>Fill GC vials with the hexane phase (upper phase) using a glass Pasteur pipette.</li>
	<li>Place the caps on the GC vials. Make sure the caps are completely sealed, and cannot be turned, to prevent evaporation from the vial.</li>
	<li>Refill the GC wash solvents (hexane), empty the waste vials, and put sample vials in auto-sampler. Before running the actual samples on the GC, first run 2 blanks containing only hexane on the GC.</li>
	<li>Run the samples on a GC-FID with a Nukol column (30 m x 0.53 mm x 1.0 &mu;m). Use inlet temperature of 250 &deg;C and use helium as carrier gas, pressure of 81.7 kPa and split ratio of 0.1:1, total flow rate: 20 ml/min. FID detector temperature of 270 &deg;C with H2 flow rate of 40 ml/min, air flow rate of 400 ml/min and He flow rate of 9.3 ml/min. Set injection volume to 1 &mu;l/sample. Pre and post wash injector with n-hexane. Initial oven temperature is set to 90 &deg;C for 0.5 min and then raised with 20 &deg;C/min until an oven temperature of 200 &deg;C is reached. Oven temperature is then maintained at 200 &deg;C for 39 min. Total run time is 45 min.</li>
</ol>

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Authors have nothing to disclose.

The described method can be used to determine the content as well as the composition of total fatty acids present in microalgal biomass. Fatty acids derived from all lipid classes, including storage (TAG) as well as membrane lipids (phospholipids and glycolipids), are detected. All fatty acid chain lengths and degrees of saturation that are present in the microalgae can be detected and distinguished. The method is based on mechanical cell disruption, solvent based lipid extraction, transesterification of fatty acids to F...

Fatty acids are one of the major constituents of microalgal biomass and typically make up between 5-50% of the cell dry weight1-3. They are mainly present in the form of glycerolipids. These glycerolipids in turn mainly consist of phospholipids, glycolipids, and triacylglycerol (TAG). Especially the fatty acids present in TAG are of commercial interest, because they can be used as a resource for production of transportation fuels, bulk chemicals, nutraceuticals (&omega;-3 fatty acids), and food commodities3-6. Microalgae can grow in sea water based cultivation media, can have a much higher areal productivity than terrestrial plants, and can be cultivated in photobioreactors at locations that are unsuitable for agriculture, possibly even offshore. For these reasons, microalgae are often considered a promising alternative to terrestrial plants for the production of biodiesel and other bulk products3-6. Potentially no agricultural land or fresh water (when cultivated in closed photobioreactors or when marine microalgae are used) is needed for their production. Therefore, biofuels derived from microalgae are considered 3rd-generation biofuels.
The total cellular content of fatty acids (% of dry weight), the lipid class composition, as well as the fatty acid length and degree of saturation are highly variable between microalgae species. Furthermore, these properties vary with cultivation conditions such as nutrient availability, temperature, pH, and light intensity1,2. For example, when exposed to nitrogen starvation, microalgae can accumulate large quantities of TAG. Under optimal growth conditions TAG typically constitutes less than 2% of dry weight, but when exposed to nitrogen starvation TAG content can increase to up to 40% of the microalgal dry weight1.
Microalgae mainly produce fatty acids with chain lengths of 16 and 18 carbon atoms, but some species can make fatty acids of up to 24 carbon atoms in length. Both saturated as well as highly unsaturated fatty acids are produced by microalgae. The latter include fatty acids with nutritional benefits (&omega;-3 fatty acids) like C20:5 (eicosapentaenoic acid; EPA) and C22:6 (docosahexaenoic acid; DHA) for which no vegetable alternatives exist1,2,4,7. The (distribution of) fatty acid chain length and degree of saturation also determines the properties and quality of algae-derived biofuels and edible oils4,8.
To develop commercial applications of microalgal derived fatty acids, reliable analytical methods for quantification of fatty acid content and composition are needed.
As also pointed out by Ryckebosch et al.9, analysis of fatty acids in microalgae distinguishes itself from other substrates (e.g. vegetable oil, food products, animal tissues etc.) because 1) microalgae are single cells surrounded by rigid cell walls, complicating lipid extraction; 2) microalgae contain a wide variety of lipid classes and the lipid class distribution is highly variable7. These different lipid classes have a wide variety in chemical structure and properties such as polarity. Also, lipid classes other than acyl lipids are produced; 3) microalgae contain a wide variety of fatty acids, ranging from 12-24 carbon atoms in length and containing both saturated as well as highly unsaturated fatty acids. Therefore, methods developed to analyze fatty acids in substrates other than microalgae, might not be suitable to analyze fatty acids in microalgae.
As reviewed by Ryckebosch et al.9, the main difference between commonly used lipid extraction procedures is in the solvent-systems that are used. Because of the large variety of lipid classes present in microalgae, each varying in polarity, the extracted lipid quantity will vary with solvents used10-12. This leads to the inconsistencies in the lipid content and composition presented in literature9,10. Depending on the solvent system used, methods based on solvent extraction without cell disruption through, for example, bead beating or sonication, might not extract all lipids because of the rigid structure of some microalgae species9,13. In the case of incomplete lipid extraction, the extraction efficiency of the different lipid classes can vary14. This can also have an effect on the measured fatty acid composition, because the fatty acid composition is variable among lipid classes7.
Our 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 in combination with a flame ionization detector (GC-FID). An internal standard in the form of a triacylglycerol (tripentadecanoin) is added prior to the analytical procedure. Possible losses during extraction and incomplete transesterification can then be corrected for. The method can be used to determine the content as well as the composition of the fatty acids present in microalgal biomass. All fatty acids present in the different acyl-lipid classes, including storage (TAG) as well as membrane lipids (glycolipids, phospholipids), are detected, identified and quantified accurately and reproducibly by this method using only small amounts of sample (5 mg). This method does not provide information about the relative abundance of different lipid classes. However, the method can be extended to separate lipid classes from each other1. The concentration and fatty acid composition of the different lipid classes can then be determined individually.
In literature several other methods are described to analyze lipids in microalgae. Some methods focus on total lipophilic components15, whereas other methods focus on total fatty acids9,16. These alternatives include gravimetric determination of total extracted lipids, direct trans-esterification of fatty acids combined with quantification using chromatography, and staining cells with lipophilic fluorescent dyes.
A commonly used alternative to quantification of fatty acids using chromatography is quantification of lipids using a gravimetric determination17,18. Advantages of a gravimetric determination are the lack of requirements for advanced and expensive equipment like a gas chromatograph; ease to set up the procedure, because of the availability of standardized analytical equipment (e.g. Soxhlet); and a gravimetric determination is less time-consuming than chromatography based methods. The major advantage of using chromatography based methods on the other hand is that in such a method only the fatty acids are measured. In a gravimetric determination the non-fatty acid containing lipids, like pigments or steroids, are also included in the determination. These non-fatty acid containing lipids can make up a large proportion (&gt;50%) of total lipids. If one is only interested in the fatty acid content (for example for biodiesel applications), it will be overestimated when a gravimetric determination is used. In addition, in a gravimetric determination the accuracy of the analytical balance used to weigh the extracted lipids determines the sample size that needs to be used. This quantity is typically much more than the amount needed when chromatography is used. Finally, another advantage of using chromatography over gravimetric determination is that chromatography provides information about the fatty acid composition.
Another alternative to our presented method is direct transesterification16,19,20. In this method lipid extraction and transesterification of fatty acids to FAMEs are combined in one step. This method is quicker than a separate extraction and transesterification step, but combining these steps limits the solvents that can be used for extraction. This might negatively influence extraction efficiency. Another advantage of a separate lipid extraction and transesterification step is that it allows for an additional lipid class separation between these steps1. This is not possible when direct transesterification is used.
Other commonly used methods to determine the lipid content in microalgae include staining the biomass with lipophilic fluorescent stains such as Nile red or BODIPY and measuring the fluorescence signal21,22. An advantage of these methods is that they are less laborious than alternative methods. A disadvantage of these methods is that the fluorescent response is, for various reasons, variable between species, cultivation conditions, lipid classes, and analytical procedures. As an example, several of these variations are caused by differences in uptake of the dye by the microalgae. Calibration using another quantitative method is therefore needed, preferably performed for all the different cultivation conditions and growth stages. Finally, this method does not provide information about the fatty acid composition and is less accurate and reproducible than chromatography based methods.
The presented method is based on the method described by Lamers et al.23 and Santos et al.24 and has also been applied by various other authors1,25-27. Also other methods are available that are based on the same principles and could provide similar results9,28.

<table border="1">
 <tbody>
 <tr>
 <td>Reagent and equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>tripentadecanoin (C15:0 TAG)</td>
 <td>Sigma Aldrich</td>
 <td>T4257</td>
 <td>CAS Number 7370-46-9</td>
 </tr>
 <tr>
 <td>TAG or FAME standards of all fatty acids expected in sample</td>
 <td>Sigma Aldrich </td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>TAG or FAME standards of all fatty acids expected in sample</td>
 <td>Lipidox</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>TAG or FAME standards of all fatty acids expected in sample</td>
 <td>Larodan</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Beadbeater</td>
 <td>Bertin Technologies</td>
 <td>Precellys 24</td>
 <td></td>
 </tr>
 <tr>
 <td>beadbeater tubes</td>
 <td>MP Biomedicals</td>
 <td>Lysing matrix E 
 116914500</td>
 <td></td>
 </tr>
 <tr>
 <td>GC-FID</td>
 <td>Hewlett-Packer</td>
 <td>HP6871</td>
 <td></td>
 </tr>
 <tr>
 <td>GC column</td>
 <td>Supelco</td>
 <td>Nukol 25357</td>
 <td></td>
 </tr>
 <tr>
 <td>Positive displacement pipette 100-1000&mu;l</td>
 <td>Mettler Toledo</td>
 <td>MR-1000</td>
 <td></td>
 </tr>
 <tr>
 <td>Positive displacement pipet tips C-1000</td>
 <td>Mettler Toledo</td>
 <td>C-1000</td>
 <td></td>
 </tr>
 <tr>
 <td>Pipetvuller, Pi-Pump 2 ml</td>
 <td>VWR</td>
 <td>612-1925</td>
 <td></td>
 </tr>
 <tr>
 <td>glass tubes</td>
 <td>VWR</td>
 <td>SCERE5100160011G1</td>
 <td></td>
 </tr>
 <tr>
 <td>TUBE 16 X 100 MM BOROSILICATE 5.1 1 * 1.000</td>
 <td>VWR</td>
 <td>SCERE5100160011G1</td>
 <td></td>
 </tr>
 <tr>
 <td>Teflon coated screw-caps</td>
 <td>VWR</td>
 <td>SCERKSSR15415BY10</td>
 <td></td>
 </tr>
 <tr>
 <td>STUART SCIENTIFIC SB2 test tube rotator</td>
 <td>VWR</td>
 <td>445-2101</td>
 <td></td>
 </tr>
 <tr>
 <td>Heated Evaporator/Concentrator</td>
 <td>Cole-Parmer</td>
 <td>YO-28690-25</td>
 <td></td>
 </tr>
 </tbody>
</table>

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 typical chromatogram that is obtained via this process is shown in Figure 1. FAMEs are separated by size and degree of saturation by the GC column and protocol used. Shorter chain length fatty acids and more saturated fatty acids (fewer double bonds) have shorter retention times. The used GC column and protocol do not intend to separate fatty acid isomers (same chain length and degree of saturation, but different positions of double bonds), but this could be achieved by using a different GC column and ...

Watch this Scientific Journal Video about Analysis of Fatty Acid Content and Composition in Microalgae at JoVE.com

Analysis of Fatty Acid Content and Composition in Microalgae

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 ...

analysis-of-fatty-acid-content-and-composition-in-microalgae

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JoVE Journal

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59.4K Views.  Wageningen University and Research Center. 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.

Video: Analysis of Fatty Acid Content and Composition in Microalgae

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 ...

Air-sampled Filter Analysis for Endotoxins and DNA Content

Outdoor aerosol research commonly uses particulate matter sampled on filters. This procedure enables various characterizations of the collected particles to be performed in parallel. The purpose of the method presented here is to obtain a highly accurate and reliable analysis of the endotoxin and DNA content of bio-aerosols extracted from filters. The extraction of high molecular weight organic molecules, such as lipopolysaccharides, from sampled filters involves shaking the sample in a pyrogen-free water-based medium. The subsequent analysis is based on an enzymatic reaction that can be detected using a turbidimetric measurement. As a result of the high organic content on the sampled filters, the extraction of DNA from the samples is performed using a commercial DNA extraction kit that was originally designed for soils and modified to improve the DNA yield. The detection and quantification of specific microbial species using quantitative polymerase chain reaction (q-PCR) analysis are described and compared with other available methods.

Two complementary analyses of atmospheric biological particles from air sampled filters are described herein: the extraction and detection of endotoxin, and of DNA.

Outdoor aerosol research commonly uses particulate matter sampled on filters. This procedure enables various characterizations of the collected particles ...

Extraction and Analysis of Microbial Phospholipid Fatty Acids in Soils

Phospholipid fatty acids (PLFAs) are key components of microbial cell membranes. The analysis of PLFAs extracted from soils can provide information about the overall structure of terrestrial microbial communities. PLFA profiling has been extensively used in a range of ecosystems as a biological index of overall soil quality, and as a quantitative indicator of soil response to land management and other environmental stressors.
The standard method presented here outlines four key steps: 1. lipid extraction from soil samples with a single-phase chloroform mixture, 2. fractionation using solid phase extraction columns to isolate phospholipids from other extracted lipids, 3. methanolysis of phospholipids to produce fatty acid methyl esters (FAMEs), and 4. FAME analysis by capillary gas chromatography using a flame ionization detector (GC-FID). Two standards are used, including 1,2-dinonadecanoyl-sn-glycero-3-phosphocholine (PC(19:0/19:0)) to assess the overall recovery of the extraction method, and methyl decanoate (MeC10:0) as an internal standard (ISTD) for the GC analysis.

Phospholipid fatty acids provide information about the structure of soil microbial communities. We present methods for extraction from soil samples with a single-phase chloroform mixture, fractionation of extracted lipids using solid phase extraction columns, and methanolysis to produce fatty acid methyl esters, which are analyzed by capillary gas chromatography.

Phospholipid fatty acids (PLFAs) are key components of microbial cell membranes. The analysis of PLFAs extracted from soils can provide information about ...

Composition and Properties of Aquafaba: Water Recovered from Commercially Canned Chickpeas

Chickpea and other pulses are commonly sold as canned products packed in a thick solution or a brine. This solution has recently been shown to produce stable foams and emulsions, and can act as a thickener. Recently interest in this product has been enhanced through the internet where it is proposed that this solution, now called aquafaba by a growing community, can be used a replacement for egg and milk protein. As aquafaba is both new and being developed by an internet based community little is known of its composition or properties. Aquafaba was recovered from 10 commercial canned chickpea products and correlations among aquafaba composition, density, viscosity and foaming properties were investigated. Proton NMR was used to characterize aquafaba composition before and after ultrafiltration through membranes with different molecular weight cut offs (MWCOs of 3, 10, or 50 kDa). A protocol for electrophoresis, and peptide mass fingerprinting is also presented. Those methods provided valuable information regarding components responsible for aquafaba functional properties. This information will allow the development of practices to produce standard commercial aquafaba products and may help consumers select products of superior or consistent utility.

Aquafaba is a viscous juice from canned chickpea that, when stirred vigorously, produces a relatively stable white froth or foam. The primary research goal is to identify the components of aquafaba that contribute viscosifying/thickening properties using nuclear magnetic resonance (NMR), ultrafiltration, electrophoresis, and peptide mass fingerprinting.

Chickpea and other pulses are commonly sold as canned products packed in a thick solution or a brine. This solution has recently been shown to produce ...

The Calibration and Use of Capacitance Sensors to Monitor Stem Water Content in Trees

Water transport and storage through the soil-plant-atmosphere continuum is critical to the terrestrial water cycle, and has become a major research focus area. Biomass capacitance plays an integral role in the avoidance of hydraulic impairment to transpiration. However, high temporal resolution measurements of dynamic changes in the hydraulic capacitance of large trees are rare. Here, we present procedures for the calibration and use of capacitance sensors, typically used to monitor soil water content, to measure the volumetric water content in trees in the field. Frequency domain reflectometry-style observations are sensitive to the density of the media being studied. Therefore, it is necessary to perform species-specific calibrations to convert from the sensor-reported values of dielectric permittivity to volumetric water content. Calibration is performed on a harvested branch or stem cut into segments that are dried or re-hydrated to produce a full range of water contents used to generate a best-fit regression with sensor observations. Sensors are inserted into calibration segments or installed in trees after pre-drilling holes to a tolerance fit using a fabricated template to ensure proper drill alignment. Special care is taken to ensure that sensor tines make good contact with the surrounding media, while allowing them to be inserted without excessive force. Volumetric water content dynamics observed via the presented methodology align with sap flow measurements recorded using thermal dissipation techniques and environmental forcing data. Biomass water content data can be used to observe the onset of water stress, drought response and recovery, and has the potential to be applied to the calibration and evaluation of new plant-level hydrodynamics models, as well as to the partitioning of remotely sensed moisture products into above- and belowground components.

The hydraulic capacitance of biomass is a key component of the vegetation water budget, which serves as a buffer against short and long-term drought stresses. Here, we present a protocol for the calibration and use of soil moisture capacitance sensors to monitor water content in the stems of large trees.

Water transport and storage through the soil-plant-atmosphere continuum is critical to the terrestrial water cycle, and has become a major research focus ...

Combining Eye-tracking Data with an Analysis of Video Content from Free-viewing a Video of a Walk in an Urban Park Environment

As individuals increasingly live in cities, methods to study their everyday movements and the data that can be collected becomes important and valuable. Eye-tracking informatics are known to connect to a range of feelings, health conditions, mental states and actions. But because vision is the result of constant eye-movements, teasing out what is important from what is noise is complex and data intensive. Furthermore, a significant challenge is controlling for what people look at compared to what is presented to them.
The following presents a methodology for combining and analyzing eye-tracking on a video of a natural and complex scene with a machine learning technique for analyzing the content of the video. In the protocol we focus on analyzing data from filmed videos, how a video can be best used to record participants&#39; eye-tracking data, and importantly how the content of the video can be analyzed and combined with the eye-tracking data. We present a brief summary of the results and a discussion of the potential of the method for further studies in complex environments.

The objective of the protocol is to detail how to collect video data for use in the laboratory; how to record eye-tracking data of participants looking at the data and how to efficiently analyze the content of the videos that they were looking at using a machine learning technique.

As individuals increasingly live in cities, methods to study their everyday movements and the data that can be collected becomes important and valuable. ...

Composition and Distribution Analysis of Bioaerosols Under Different Environmental Conditions

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. In this study, we described a protocol for multiple analyses of the biological compositions in environmental PM. Five experiments are presented: (1) PM number monitoring by using a laser particle counter; (2) PM collection by using a cyclonic aerosol sampler; (3) PM collection by using a high-volume air sampler with filters; (4) culturable microbes collection by the Andersen six-stage sampler; and (5) detection of biological composition of environmental PM by bacterial 16SrDNA and fungal ITS region sequencing. We selected hazy days and a livestock farm as two typical examples of the application in this protocol. In this study, these two sampling methods, cyclonic aerosol sampler and filter sampler, showed different sampling efficiency. The cyclonic aerosol sampler performed much better in terms of collecting bacteria, while these two methods showed the same efficiency in collecting fungi. Filter samplers can work under low temperature conditions while cyclonic aerosol samplers have a sampling limitation for temperature. A solid impacting sampler, such as an Andersen six-stage sampler, can be used to sample bioaerosols directly into the culture medium, which increases the survival rate of culturable microorganisms. However, this method mainly relies on culture while more than 99% of microbes cannot be cultured. DNA extracted from the culturable bacteria collected by the Andersen six-stage sampler and samples collected by cyclonic aerosol sampler and filter sampler were detected by bacterial 16S rDNA and fungal ITS region sequencing.All the methods above may have wide application in many fields of study, such as environmental monitoring and airborne pathogen detection. From these results, we can conclude that these methods can be used under different conditions and may help other researchers further explore the health impacts of environmental bioaerosols.

Understanding the biological composition of environmental particulate matter is important for the study of its significant impacts on human health and disease spread. Here, we used three types of bioaerosol sampling methods and a biological analysis of airborne microbes to better explore airborne microbial communities under different environmental conditions.

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. 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 ...

Manufacturing Simple and Inexpensive Soil Surface Temperature and Gravimetric Water Content Sensors

Quantifying temperature and moisture at the soil surface is essential for understanding how soil surface biota respond to changes in the environment. However, at the soil surface these variables are highly dynamic and standard sensors do not explicitly measure temperature or moisture in the upper few millimeters of the soil profile. This paper describes methods for manufacturing simple, inexpensive sensors that simultaneously measure the temperature and moisture of the upper 5 mm of the soil surface. In addition to sensor construction, steps for quality control, as well as for calibration for various substrates, are explained. The sensors incorporate a Type E thermocouple to measure temperature and assess soil moisture by measuring the resistance between two gold-plated metal probes at the end of the sensor at a depth of 5 mm. The methods presented here can be altered to customize probes for different depths or substrates. These sensors have been effective in a variety of environments and have endured months of heavy rains in tropical forests as well as intense solar radiation in deserts of the southwestern U.S. Results demonstrate the effectiveness of these sensors for evaluating warming, drying, and freezing of the soil surface in a global change experiment.

Accurately measuring temperature and water content of the upper 5 mm of the soil surface can improve our understanding of environmental controls on biological, chemical, and physical processes. Here we describe a protocol for manufacturing, calibrating, and conducting measurements with soil surface temperature and moisture sensors.

Quantifying temperature and moisture at the soil surface is essential for understanding how soil surface biota respond to changes in the environment. ...