This research was funded by PATROLS &#8211; Advanced Tools for NanoSafety Testing, Grant agreement 760813 under Horizon 2020 research and innovation program.

1. Description of the LEVITATT setup
<ol>
	<li>Use 20 mL scintillation glass vials (Figure 1, insert 1) allowing light penetration. Alternatively, light penetrable plastic vials can be used. Quantify the light intensity using a photometer.</li>
	<li>Use at least a 4 mL test suspension at the beginning of the test to allow for quantification of biomass and for characterization/quantification of nanomaterials during and after incubation (Figure 1, insert 2).</li>
	<li>Fit the 20 mL scintillation vials with a cap (Figure 1, insert 3) where a small hole is drilled (approximately 1 mm in diameter) to allow for CO2 exchange with the atmosphere. This exchange is crucial to ensure stable pH and CO2 levels during testing.</li>
	<li>For volatile substances, use an air-tight Teflon coated cap to allow for CO2 enriching of the headspace using a syringe9 or completely closed flasks with no gas phase in which CO2 is maintained in solution by an enriched sodium bicarbonate (NaHCO3) buffer system10.</li>
	<li>Fasten the vials with clamps mounted on the exterior casing (Figure 1, insert 4).</li>
	<li>Use an LED light source located below the test vials (Figure 1, insert 5) providing a uniform fluorescent illumination of &#8220;cool-white&#8221; or &#8220;daylight&#8221; type and a light intensity in the range 60&#8211;120 &#181;E&#8729;m-2&#8729;s-1 measured in the photosynthetically effective wavelength range of 400 nm to 700 nm. The setup employs adjustable light intensity in the range 5&#8211;160 &#181;E&#8729;m-2&#8729;s-1 by fitting a light dimmer to the source. This allows for testing at higher and lower light intensities.</li>
	<li>Mount the setup on an orbital shaker to agitate samples throughout the duration of the test. This keeps the cells in free suspension and facilitates CO2 mass transfer from air to water (Figure 1, insert 6).</li>
	<li>Place the setup in a temperature-controlled room or a thermostatic cabinet to maintain stable temperatures throughout testing (Figure 1, insert 7).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61209/61209fig1v2.jpg" /> 
Figure 1: Picture of LED Vertical Illumination Table for Algal Toxicity Tests (LEVITATT).&#160;1) 20 mL glass scintillation vials for incubation, 2) 4 mL sample for analysis, 3) lid with drilled hole for CO2 exchange, 4) casing for defined light conditions, 5) LED light source located in the center of the casing, 6) orbital shaker for agitation during the experiment, and 7) a thermostatic cabinet. <a href="https://www.jove.com/files/ftp_upload/61209/61209fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Preparation of algal growth medium
<ol>
	<li>The ISO 8692 algal growth medium consists of four different stock solutions. Weigh out the appropriate amount of salts and dilute in ultrapure water according to Table 1.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Stock solutions</td>
			<td>Nutrient</td>
			<td>Concentration in stock solution</td>
			<td>Concentration in test solution</td>
		</tr>
		<tr>
			<td rowspan="5">1: Macronutrients</td>
			<td>NH4Cl</td>
			<td>1.5 g/L</td>
			<td>15 mg/L (N: 3.9 mg/L)</td>
		</tr>
		<tr>
			<td>MgCl2&#8729;6H2O</td>
			<td>1.2 g/L</td>
			<td>12 mg/L (Mg: 2.9 mg/L)</td>
		</tr>
		<tr>
			<td>CaCl2&#8729;2H2O</td>
			<td>1.8 g/L</td>
			<td>18 mg/L (Ca: 4.9 mg/L)</td>
		</tr>
		<tr>
			<td>MgSO4&#8729;7H2O</td>
			<td>1.5 g/L</td>
			<td>15 mg/L (S: 1.95 mg/L)</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>0.16 g/L</td>
			<td>1.6 mg/L (P: 0,36 mg/L)</td>
		</tr>
		<tr>
			<td rowspan="2">2: Fe-EDTA</td>
			<td>FeCl3&#8729;6H2O</td>
			<td>64 mg/L</td>
			<td>64 &#181;g/L (Fe: 13 &#181;g/L)</td>
		</tr>
		<tr>
			<td>Na2EDTA&#8729;2H2O</td>
			<td>100 mg/L</td>
			<td>100 &#181;g/L</td>
		</tr>
		<tr>
			<td rowspan="6">3: Trace elements</td>
			<td>H3BO3a</td>
			<td>185 mg/L</td>
			<td>185 &#181;g/L (B: 32 &#181;g/L)</td>
		</tr>
		<tr>
			<td>MnCl2&#8729;4H2O</td>
			<td>415 mg/L</td>
			<td>415 &#181;g/L (Mn: 115 &#181;g/L)</td>
		</tr>
		<tr>
			<td>ZnCl2</td>
			<td>3 mg/L</td>
			<td>3 &#181;g/L (Zn: 1.4 &#181;g/L)</td>
		</tr>
		<tr>
			<td>CoCl2&#8729;6H2O</td>
			<td>1.5 mg/L</td>
			<td>1.5 &#181;g/L (Co: 0.37 &#181;g/L)</td>
		</tr>
		<tr>
			<td>CuCl2&#8729;2H2O</td>
			<td>0.01 mg/L</td>
			<td>0.01 &#181;g/L (Cu: 3.7 ng/L)</td>
		</tr>
		<tr>
			<td>Na2MoO4&#8729;2H2O</td>
			<td>7 mg/L</td>
			<td>7 &#181;g/L (Mo: 2.8 &#181;g/L)</td>
		</tr>
		<tr>
			<td>4: NaHCO3</td>
			<td>NaHCO3</td>
			<td>50 g/L</td>
			<td>50 mg/L (C: 7.14 mg/L)</td>
		</tr>
	</tbody>
</table>
Table 1: Concentrations of nutrients in stock solutions for algal growth medium
NOTE: H3BO3 can be dissolved by adding 0.1 M NaOH. EDTA should be removed when testing metals, to avoid complexation with metal ions. Sterilize the stock solutions by membrane filtration (mean pore diameter 0.2 &#181;m) or by autoclaving (120 &#176;C, 15 min). Do no autoclave stock solutions 2 and 4, but sterilize them by membrane filtration. Store the solutions in the dark at 4 &#176;C.
<ol start="2">
	<li>To produce 1 L of algal growth medium, transfer 500 mL sterilized ultrapure water into a 1 L sterilized volumetric flask and add 10 mL of stock solution 1: Macronutrients, 1 mL of stock solution 2: Fe-EDTA, 1 mL of stock solution 3: Trace elements, and 1 mL of stock solution 4: NaHCO3.</li>
	<li>Fill up to 1 L with sterilized ultrapure water, stopper the flask and shake thoroughly to homogenize the algal growth medium.</li>
	<li>Equilibrate the solution before use by leaving it overnight in contact with air or by bubbling with sterile, filtered air for 30 min. After equilibration, adjust the pH, if necessary, to pH 8.1 &#177; 0.2, with either 1 M HCl or 1 M NaOH.</li>
</ol>
3. Setting up the algal test
NOTE: A flow diagram of the algal test procedure is shown in Figure 2.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61209/61209fig2v6.jpg" /> 
Figure 2: Flow diagram of the algal test setup. <a href="https://www.jove.com/files/ftp_upload/61209/61209fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Prepare a stock solution of the test compound at the desired highest test concentration in the algal growth medium prepared according to step 2. For preparation of stock solutions/suspensions, follow OECD 201 (for soluble compounds) or OECD 318 (for nanomaterials).</li>
	<li>Measure the pH in the stock solution. If it deviates more than one unit from the algal growth medium, adjust the pH to 8 with either 1 M HCl or 1 M NaOH.</li>
	<li>Calculate the inoculum volume needed to reach a final cell concentration of 1 x 104 cells/mL in a 25 mL test solution. 
	NOTE: The inoculum should come from a culture of uncontaminated exponentially growing Raphidocelis subcapitata grown using the LEVITATT setup.</li>
	<li>Calculate the amount of stock solution to add to each 25 mL volumetric flask to obtain the desired test concentrations. The factor between each concentration should not exceed 3.2.</li>
	<li>Mark one 25 mL volumetric flask for each chosen concentration and an additional 25 mL volumetric flask marked control.</li>
	<li>Add the amount of stock solution of the test compound needed to reach the desired concentrations to the 25 mL volumetric flask. Do not add stock solution to the control.</li>
	<li>Add the medium to each 25 mL volumetric flask to reach a volume of approximately 20 mL.</li>
	<li>Add the volume of inoculum calculated in step 3.3 to each 25 mL volumetric flask. Add the medium to each 25 mL volumetric flask to a final total volume of 25 mL.</li>
	<li>Stopper the flasks and mix thoroughly by turning the flasks two times vertically.</li>
	<li>Transfer 0.4 mL from each flask into individual screw cap vials and add 1.6 mL of acetone (saturated with MgCO3): one sample for each test concentration and the control. Close the lids tightly and store in the dark at room temperature until fluorescence measurements (section 4).</li>
	<li>Pipet 4 mL of each test solution into 20 mL scintillation vials (3 replicates per concentration and 5 replicates for the control). Screw lids on the scintillation vials. Remember that the lids must have a drilled hole (approximately 1 mm in diameter) to allow for CO2 exchange.</li>
	<li>After 24 h, 48 h, and 72 h, pipet 0.4 mL from each vial into screw cap vials and add 1.6 mL of acetone (saturated with MgCO3). Close the lids tightly and store in the dark at room temperature until fluorescence measurements (section 4).</li>
	<li>After the last sample is taken at 72 h, gently pool the three replicates for a given concentration in one vial and measure the pH. Repeat for all concentrations and the control. The pH should not deviate more than 1.5 units from the initial pH for any of the samples measured.</li>
	<li>Discharge the remaining liquids into a waste container following your institutional rules and regulations.</li>
</ol>
4. Analyzing algal test samples
<ol>
	<li>Use a fluorescence spectrophotometer to measure the algal biomass (here expressed as chlorophyll A). The peak emission for chlorophyll A is 420 nm for the excitation wavelength and 671 nm for the emission wavelength.</li>
	<li>Measure the fluorescence of each individual sample three times and calculate the average value for each sample.</li>
	<li>Use equation 1 to calculate the growth rate. The measured fluorescence (relative units) can be used directly as the biomass parameter in equation 1. 
	Equation 1: &#181; = (ln Nt &#8211; ln N0) / t 
	where &#181; is the growth rate (d-1), N0 is the initial biomass, Nt is the biomass at time t, and t is the length of the test period (d). Note, N0 and Nt should be expressed in the same unit.</li>
	<li>Use a statistical software to fit a non-linear regression curve (e.g., a log-logistic or Weibull function) to the growth rate data to obtain effective concentration values at 10%, 20%, and 50% inhibition. In the supplementary information an example of code for fitting in the statistical software R using the DRC package11 is given.</li>
</ol>

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The authors have nothing to disclose.

Phytoplankton converts solar energy and carbon dioxide to organic matter and thus holds a pivotal role in the aquatic ecosystem. For this reason, algal growth rate inhibition tests are included as one of three mandatory aquatic toxicity tests required for regulatory risk assessment of chemicals. The ability to perform a reliable and reproducible algal toxicity test is key in this regard. Test setups using Erlenmeyer flasks introduces a range of variabilities and inconveniences as described in the introduction. To circumv...

The algal toxicity test is one of only three mandatory tests used to generate the ecotoxicity data required for pre- and post-market registration of chemicals by European and international regulations (e.g., REACH1 and TSCA (USA)). For this purpose, standardized algal test guidelines have been developed by international organizations (e.g., ISO and OECD). These testing standards and guidelines prescribe ideal test conditions in terms of pH, temperature, carbon dioxide levels and light intensity. However, maintaining stable test conditions during algal testing is in practice difficult and the results suffer from problems with reproducibility and reliability for a range of chemical substances and nanomaterials (often referred to as &#8220;difficult substances&#8221;)2. Most of the existing algal toxicity testing setups operate with relatively large volumes (100&#8211;250 mL) situated on an orbital shaker inside an incubator. Such a setup limits the number of test concentrations and replicates achievable and high volumes of algal culture and test material. Additionally, these setups rarely have a uniform light field and reliable lighting conditions are furthermore difficult to obtain in large flasks, partly as light intensity decreases exponentially the further the light travels and partly due to the flask geometry. Alternative setups comprise plastic microtiter3 plates containing small sample volumes that do not allow for adequate sampling volumes to measure pH, additional biomass measurements, pigment extraction or other analyses requiring destructive sampling. One particular challenge using existing setups for algal toxicity testing of nanomaterials and substances forming colored suspensions is the interference or blocking of the light available to the algal cells, often referred to as &#8220;shading&#8221;4,5. Shading may occur within vials by the test material and/or interactions between the test material and the algal cells, or shading can occur between vials, due to their positioning relative to each other and the light source.
The method is based on the small-scale algal toxicity test setup introduced by Arensberg et al.6 that allows for testing in compliance with standards such as OECD 2017, and ISO 86928. The method is further optimized to address the limitations stated above by: 1) utilizing the LED light technology to ensure uniform light conditions with minimal heat generation, 2) providing adequate sample volume for chemical/biological analysis while maintaining constant pH, CO2 levels, and 3) enabling the use of versatile test container material for testing of volatile substances or substances with a high sorption potential.

<table>
	<tbody>
		<tr>
			<td>Acetone</td>
			<td>Sigma-Aldrich</td>
			<td>V179124</td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>254134</td>
			<td></td>
		</tr>
		<tr>
			<td>BlueCap bottles (1L)</td>
			<td>Buch &#38; Holm A/S</td>
			<td>&#160;9072335</td>
			<td></td>
		</tr>
		<tr>
			<td>Boric acid</td>
			<td>Sigma-Aldrich</td>
			<td>B0394</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>208290</td>
			<td></td>
		</tr>
		<tr>
			<td>Clear acrylic sheet (40x40 cm)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cobalt(II) chloride hexahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>255599</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper(II) chloride dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>307483</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid disodium salt dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>&#160;E5134</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence Spectrophotometer F-7000</td>
			<td>Hitachi</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric acid</td>
			<td>Sigma-Aldrich</td>
			<td>258148</td>
			<td></td>
		</tr>
		<tr>
			<td>Iron(III) chloride hexahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>236489</td>
			<td></td>
		</tr>
		<tr>
			<td>LED light source</td>
			<td>Helmholt Elektronik A/S</td>
			<td>H35161</td>
			<td>Neutral White, 6500K</td>
		</tr>
		<tr>
			<td>Magnesium chloride hexahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>M9272</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>230391</td>
			<td></td>
		</tr>
		<tr>
			<td>Manganese(II) chloride tetrahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>221279</td>
			<td></td>
		</tr>
		<tr>
			<td>Orbital shaker</td>
			<td>IKA</td>
			<td>2980200</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic</td>
			<td>Sigma-Aldrich</td>
			<td>P0662</td>
			<td></td>
		</tr>
		<tr>
			<td>Raphidocelis subcapitata</td>
			<td>NORCCA</td>
			<td></td>
			<td>NIVA-CHL1 strain</td>
		</tr>
		<tr>
			<td>Scintillation vials (20 mL)</td>
			<td>Fisherscientific</td>
			<td>11526325</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>S6014</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Sigma-Aldrich</td>
			<td>415413</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium molybdate dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>331058&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring clamp</td>
			<td>Frederiksen Scientific A/S</td>
			<td>472002</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermostatic cabinet</td>
			<td>VWR</td>
			<td>WTWA208450</td>
			<td>Alternative: temperature controlled room</td>
		</tr>
		<tr>
			<td>Ventilation pipe (&#216;125 mm)</td>
			<td>Silvan</td>
			<td>22605630165</td>
			<td></td>
		</tr>
		<tr>
			<td>Volumetric flasks (25 mL)</td>
			<td>DWK Life Sciences</td>
			<td>246781455</td>
			<td></td>
		</tr>
		<tr>
			<td>Zinc chloride</td>
			<td>Sigma-Aldrich</td>
			<td>208086</td>
			<td></td>
		</tr>
	</tbody>
</table>

a small-scale setup for algal toxicity testing of nanomaterials and other difficult substances

Ecotoxicity data is a requirement for pre- and post-market registration of chemicals by European and international regulations (e.g., REACH). The algal toxicity test is frequently used in regulatory risk assessment of chemicals. In order to achieve high reliability and reproducibility the development of standardized guidelines is vital. For algal toxicity testing, the guidelines require stable and uniform conditions of parameters such as pH, temperature, carbon dioxide levels and light intensity. Nanomaterials and other so-called difficult substances can interfere with light causing a large variation in results obtained hampering their regulatory acceptance. To address these challenges, we have developed LEVITATT (LED Vertical Illumination Table for Algal Toxicity Tests). The setup utilizes LED illumination from below allowing for a homogenous light distribution and temperature control while also minimizing intra-sample shading. The setup optimizes the sample volume for biomass quantification and does at the same time ensure a sufficient influx of CO2 to support exponential growth of the algae. Additionally, the material of the test containers can be tailored to minimize adsorption and volatilization. When testing colored substances or particle suspensions, the use of LED lights also allows for increasing the light intensity without additional heat generation. The compact design and minimal equipment requirements increase the possibilities for implementation of the LEVITATT in a wide range of laboratories. While compliant with standardized ISO and OECD guidelines for algal toxicity testing, LEVITATT also showed a lower inter-sample variability for two reference substances (3,5-Dicholorophenol and K2Cr2O7) and three nanomaterials (ZnO, CeO2, and BaSO4) compared to Erlenmeyer flasks and microtiter plates.

An initial test with a reference substance is carried out to determine the sensitivity of the algal strain. Reference substances regularly used for R. subcapitata are potassium dichromate and 3,5-Dichlorphenol7,8. Figure 3 and Table 2 show a representative result of an algal test including curve fitting and statistical outputs when the DRC package in R is applied to the growth rates.
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Watch this Scientific Journal Video about A Small-Scale Setup for Algal Toxicity Testing of Nanomaterials and Other Difficult Substances at JoVE.com

A Small-Scale Setup for Algal Toxicity Testing of Nanomaterials and Other Difficult Substances

We demonstrate algal toxicity testing for difficult substances (e.g., colored substances or nanomaterials) using a setup illuminated vertically with an LED.

Ecotoxicity data is a requirement for pre- and post-market registration of chemicals by European and international regulations (e.g., REACH). The algal ...

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JoVE Journal

Environment

5.6K Views.  Technical University of Denmark. We demonstrate algal toxicity testing for difficult substances (e.g., colored substances or nanomaterials) using a setup illuminated vertically with an LED.

Video: A Small-Scale Setup for Algal Toxicity Testing of Nanomaterials and Other Difficult Substances

Preparation and Testing of Impedance-based Fluidic Biochips with RTgill-W1 Cells for Rapid Evaluation of Drinking Water Samples for Toxicity

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial (RTgill-W1) cells for use in a field portable water toxicity sensor. A monolayer of RTgill-W1 cells forms on the sensing electrodes enclosed within the biochips. The biochips are then used for testing in a field portable electric cell-substrate impedance sensing (ECIS) device designed for rapid toxicity testing of drinking water. The manuscript further describes how to run a toxicity test using the prepared biochips. A control water sample and the test water sample are mixed with pre-measured powdered media and injected into separate channels of the biochip. Impedance readings from the sensing electrodes in each of the biochip channels are measured and compared by an automated statistical software program. The screen on the ECIS instrument will indicate either "Contamination Detected" or "No Contamination Detected" within an hour of sample injection. Advantages are ease of use and rapid response to a broad spectrum of inorganic and organic chemicals at concentrations that are relevant to human health concerns, as well as the long-term stability of stored biochips in a ready state for testing. Limitations are the requirement for cold storage of the biochips and limited sensitivity to cholinesterase-inhibiting pesticides. Applications for this toxicity detector are for rapid field-portable testing of drinking water supplies by Army Preventative Medicine personnel or for use at municipal water treatment facilities.

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial cells for use in a field portable electric cell-substrate impedance sensor. The protocol for running a rapid drinking water toxicity test with the sensor is also described.

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial (RTgill-W1) cells for use in a field portable water toxicity ...

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

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

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

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

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

Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

This protocol provides multiple methods for the analysis and quantification of predatory feeding behaviors in nematodes. Many nematode species including Pristionchus pacificus display complex behaviors, the most striking of which is the predation of other nematode larvae. However, as these behaviors are absent in the model organism Caenorhabditis elegans, they have thus far only recently been described in detail along with the development of reliable behavioral assays 1. These predatory behaviors are dependent upon phenotypically plastic but fixed mouth morphs making the correct identification and categorization of these animals essential. In P. pacificus there are two mouth types, the stenostomatous and eurystomatous morphs 2, with only the wide mouthed eurystomatous containing an extra tooth and being capable of killing other nematode larvae. Through the isolation of an abundance of size matched prey larvae and subsequent exposure to predatory nematodes, assays including both "corpse assays" and "bite assays" on correctly identified mouth morph nematodes are possible. These assays provide a means to rapidly quantify predation success rates and provide a detailed behavioral analysis of individual nematodes engaged in predatory feeding activities. In addition, with the use of a high-speed camera, visualization of changes in pharyngeal activity including tooth and pumping dynamics are also possible.

Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

Behavioral assays provide powerful tools for understanding neuronal function. Here we present several protocols for quantifying predatory feeding behavior found in the model nematode Pristionchus pacificus and its relatives. Additionally, we provide methods for analyzing predatory feeding adaptations including mouth structures and teeth.

This protocol provides multiple methods for the analysis and quantification of predatory feeding behaviors in nematodes. Many nematode species including ...

Extraction of Structural Extracellular Polymeric Substances from Aerobic Granular Sludge

To evaluate and develop methodologies for the extraction of gel-forming extracellular polymeric substances (EPS), EPS from aerobic granular sludge (AGS) was extracted using six different methods (centrifugation, sonication, ethylenediaminetetraacetic acid (EDTA), formamide with sodium hydroxide (NaOH), formaldehyde with NaOH and sodium carbonate (Na2CO3) with heat and constant mixing). AGS was collected from a pilot wastewater treatment reactor. The ionic gel-forming property of the extracted EPS of the six different extraction methods was tested with calcium ions (Ca2+). From the six extraction methods used, only the Na2CO3 extraction could solubilize the hydrogel matrix of AGS. The alginate-like extracellular polymers (ALE) recovered with this method formed ionic gel beads with Ca2+. The Ca2+-ALE beads were stable in EDTA, formamide with NaOH and formaldehyde with NaOH, indicating that ALE are one part of the structural polymers in EPS. It is recommended to use an extraction method that combines physical and chemical treatment to solubilize AGS and extract structural EPS.

The protocol provides a methodology to solubilize aerobic granular sludge in order to extract alginate-like extracellular polymers (ALE).

To evaluate and develop methodologies for the extraction of gel-forming extracellular polymeric substances (EPS), EPS from aerobic granular sludge (AGS) ...

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, ...

Functionalization and Dispersion of Carbon Nanomaterials Using an Environmentally Friendly Ultrasonicated Ozonolysis Process

Functionalization of carbon nanomaterials is often a critical step that facilitates their integration into larger material systems and devices. In the as-received form, carbon nanomaterials, such as carbon nanotubes (CNTs) or graphene nanoplatelets (GNPs), may contain large agglomerates. Both agglomerates and impurities will diminish the benefits of the unique electrical and mechanical properties offered when CNTs or GNPs are incorporated into polymers or composite material systems. Whilst a variety of methods exist to functionalize carbon nanomaterials and to create stable dispersions, many the processes use harsh chemicals, organic solvents, or surfactants, which are environmentally unfriendly and may increase the processing burden when isolating the nanomaterials for subsequent use. The current research details the use of an alternative, environmentally friendly technique for functionalizing CNTs and GNPs. It produces stable, aqueous dispersions free of harmful chemicals. Both CNTs and GNPs can be added to water at concentrations up to 5 g/L and can be recirculated through a high-powered ultrasonic cell. The simultaneous injection of ozone into the cell progressively oxidizes the carbon nanomaterials, and the combined ultrasonication breaks down agglomerates and immediately exposes fresh material for functionalization. The prepared dispersions are ideally suited for the deposition of thin films onto solid substrates using electrophoretic deposition (EPD). CNTs and GNPs from the aqueous dispersions can be readily used to coat carbon- and glass-reinforcing fibers using EPD for the preparation of hierarchical composite materials.

Here, a novel method for the functionalization and stable dispersion of carbon nanomaterials in aqueous environments is described. Ozone is injected directly into an aqueous dispersion of carbon nanomaterial that is continuously recirculated through a high-powered ultrasonic cell.

Functionalization of carbon nanomaterials is often a critical step that facilitates their integration into larger material systems and devices. In the ...

Dispersion of Nanomaterials in Aqueous Media: Towards Protocol Optimization

The sonication process is commonly used for de-agglomerating and dispersing nanomaterials in aqueous based media, necessary to improve homogeneity and stability of the suspension. In this study, a systematic step-wise approach is carried out to identify optimal sonication conditions in order to achieve a stable dispersion. This approach has been adopted and shown to be suitable for several nanomaterials (cerium oxide, zinc oxide, and carbon nanotubes) dispersed in deionized (DI) water. However, with any change in either the nanomaterial type or dispersing medium, there needs to be optimization of the basic protocol by adjusting various factors such as sonication time, power, and sonicator type as well as temperature rise during the process. The approach records the dispersion process in detail. This is necessary to identify the time points as well as other above-mentioned conditions during the sonication process in which there may be undesirable changes, such as damage to the particle surface thus affecting surface properties. Our goal is to offer a harmonized approach that can control the quality of the final, produced dispersion. Such a guideline is instrumental in ensuring dispersion quality repeatability in the nanoscience community, particularly in the field of nanotoxicology.

Here, we present a step-wise protocol for the dispersion of nanomaterials in aqueous media with real-time characterization to identify the optimal sonication conditions, intensity, and duration for improved stability and uniformity of nanoparticle dispersions without impacting the sample integrity.

The sonication process is commonly used for de-agglomerating and dispersing nanomaterials in aqueous based media, necessary to improve homogeneity and ...

Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, and the majority of extant heteropterans retain this trophic strategy. Some heteropterans have transitioned to feeding on vertebrate blood (such as the kissing bugs, Triatominae; and bed bugs, Cimicidae) while others have reverted to feeding on plants (most Pentatomomorpha). However, with the exception of saliva used by kissing bugs to facilitate blood-feeding, little is known about heteropteran venoms compared to the venoms of spiders, scorpions and snakes.
One obstacle to the characterization of heteropteran venom toxins is the structure and function of the venom/labial glands, which are both morphologically complex and perform multiple biological roles (defense, prey capture, and extra-oral digestion). In this article, we describe three methods we have successfully used to collect heteropteran venoms. First, we present electrostimulation as a convenient way to collect venom that is often lethal when injected into prey animals, and which obviates contamination by glandular tissue. Second, we show that gentle harassment of animals is sufficient to produce venom extrusion from the proboscis and/or venom spitting in some groups of heteropterans. Third, we describe methods to harvest venom toxins by dissection of anaesthetized animals to obtain the venom glands. This method is complementary to other methods, as it may allow harvesting of toxins from taxa in which electrostimulation and harassment are ineffective. These protocols will enable researchers to harvest toxins from heteropteran insects for structure-function characterization and possible applications in medicine and agriculture.

Although many insects in the suborder Heteroptera (Insecta: Hemiptera) are venomous, their venom composition and the functions of their venom toxins are mostly unknown. This protocol describes methods to harvest heteropteran venoms for further characterization, using electrostimulation, harassment, and gland dissection.

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, ...

A High-throughput Assay for the Prediction of Chemical Toxicity by Automated Phenotypic Profiling of Caenorhabditis elegans

Applying toxicity testing of chemicals in higher order organisms, such as mice or rats, is time-consuming and expensive, due to their long lifespan and maintenance issues. On the contrary, the nematode Caenorhabditis elegans (C. elegans) has advantages to make it an ideal choice for toxicity testing: a short lifespan, easy cultivation, and efficient reproduction. Here, we describe a protocol for the automatic phenotypic profiling of C. elegans in a 384-well plate. The nematode worms are cultured in a 384-well plate with liquid medium and chemical treatment, and videos are taken of each well to quantify the chemical influence on 33 worm features. Experimental results demonstrate that the quantified phenotype features can classify and predict the acute toxicity for different chemical compounds and establish a priority list for further traditional chemical toxicity assessment tests in a rodent model.

A High-throughput Assay for the Prediction of Chemical Toxicity by Automated Phenotypic Profiling of Caenorhabditis elegans

A quantitative method has been developed to identify and predict the acute toxicity of chemicals by automatically analyzing the phenotypic profiling of Caenorhabditis elegans. This protocol describes how to treat worms with chemicals in a 384-well plate, capture videos, and quantify toxicological related phenotypes.

Applying toxicity testing of chemicals in higher order organisms, such as mice or rats, is time-consuming and expensive, due to their long lifespan and ...

Development and Testing of Species-specific Quantitative PCR Assays for Environmental DNA Applications

New, non-invasive methods for detecting and monitoring species presence are being developed to aid in fisheries and wildlife conservation management. The use of environmental DNA (eDNA) samples for detecting macrobiota is one such group of methods that is rapidly becoming popular and being implemented in national management programs. Here we focus on the development of species-specific targeted assays for probe-based quantitative PCR (qPCR) applications. Using probe-based qPCR offers greater specificity than is possible with primers alone. Furthermore, the ability to quantify the amount of DNA in a sample can be useful in our understanding of the ecology of eDNA and the interpretation of eDNA detection patterns in the field. Careful consideration is needed in the development and testing of these assays to ensure the sensitivity and specificity of detecting the target species from an environmental sample. In this protocol we will delineate the steps needed to design and test probe-based assays for the detection of a target species; including creation of sequence databases, assay design, assay selection and optimization, testing assay performance, and field validation. Following these steps will help achieve an efficient, sensitive, and specific assay that can be used with confidence. We demonstrate this process with our assay designed for populations of the mucket (Actinonaias ligamentina), a freshwater mussel species found in the Clinch River, USA.

Environmental DNA assays require rigorous design, testing, optimization and validation before the collection of field data can begin. Here, we present a protocol to take users through each step of designing a species-specific, probe-based qPCR assay for the detection and quantification of a target species DNA from environmental samples.

New, non-invasive methods for detecting and monitoring species presence are being developed to aid in fisheries and wildlife conservation management. The ...