The authors acknowledge support from SNSF PRIMA grant 179834 (to E.S.), discretionary funding from ETH (to R.S.), ETH Zurich Research Grant (to R.S. and J.J.M.), and discretionary funding from Eawag (to J.J.M.). The authors would like to thank Roberto Pioli for illustrating the experimental setup in Figure 1B and Ela Burmeister for the silicon wafer preparation.

1. Silicon wafer preparation
<ol>
	<li>Design the geometries of the microfluidic channel in computer-aided design (CAD; see Table of Materials) software and print it onto a transparent film to create the photomask (Figure 1A).</li>
	<li>Fabricate the master mold by soft lithography (under clean-room conditions) following the steps below.
	<ol>
		<li>Bake the silicon wafer at 200 &#176;C for 2 h.</li>
		<li>Place the wafer at the center of a spin-coater and pour SU8 3050 photoresist (see Table of Materials) onto the wafer. Spin coat at 1,700 rpm for 40 s with a 10 s/100 rpm ramp time. 
		NOTE: The spin-coating parameters were set to obtain a target thickness of 100 &#181;m for the SU8 3050.</li>
		<li>After the spin-coating process, soft bake the silicon wafer at 65 &#176;C for 600 s and 95 &#176;C for 2,700 s. Let the wafer cool at room temperature overnight. 
		NOTE: The overnight cooling enhances the adhesion of the SU8 to the wafer.</li>
		<li>Place the photomask (step 1.1) onto the wafer and expose it to UV light, with an exposure energy of 250 mJ/cm2 and at a wavelength of 350 nm.</li>
		<li>Post-exposure, bake the exposed substrate at 65 &#176;C for 60 s and 95 &#176;C for 300 s.</li>
		<li>Develop the silicon wafer to obtain the master mold by immersing it in a beaker filled with mrDev600 developer (see Table of Materials). Gently shake the beaker for 1,800 s to wash out the unpolymerized resist. Then, splash wash by spraying isopropanol on the silicon wafer and air dry.</li>
		<li>Hard bake the silicon wafer at 200 &#176;C for 1,800 s.</li>
	</ol>
	</li>
	<li>Silanize the master mold through vapor deposition of 20 &#181;L of Trichloro (1H, 1H, 2H, 2H-perfluorooctyl) silane (see Table of Materials) placed on a glass slide next to the mold for 40 min in a vacuum desiccator, creating a gauge pressure of 100 mbar.</li>
</ol>
2. Fabrication of the microfluidic device
NOTE: The fabrication procedure described here is for a microfluidic device with one microfluidic channel. However, the same method can be applied to fabricate a microfluidic device with multiple microfluidic channels in parallel.
<ol>
	<li>Mix the elastomer with its crosslinker at a ratio of 10:1 (see Table of Materials) to prepare a PDMS mixture. Stir the mixture until it gets uniformly mixed and turns opaque due to the enclosed air bubbles.</li>
	<li>Degas the mixture in a vacuum desiccator, creating a gauge pressure of 100 mbar until the entrapped air bubbles are removed and it looks transparent. The time required for degassing is typically 30 min.</li>
	<li>Place the master mold (step 1) in a cell culture dish (see Table of Materials). Pour 20 g of the PDMS mixture on the master mold to produce channels with a final thickness of 5 mm.</li>
	<li>Bake the master mold at 70 &#176;C for 2 h.</li>
	<li>Cut the cured PDMS around the microfluidic channel (at a distance of approximately 3 mm) using a blade, and then peel the PDMS microfluidic channel off the master mold.</li>
	<li>To create the microfluidic channels&#39; inlet and outlet, punch holes with a biopsy punch (diameter of 1.5 mm) at its extremities (top of the triangles, see Figure 1A). Punch one additional hole at the center of the inlet triangle to install the pressure sensor later.</li>
	<li>Wash a glass slide and the microfluidic channel with a commercially available 1% detergent solution (see Table of Materials) for 5 min, then rinse them with deionized water. Thereafter, wash the PDMS microfluidic channel and the glass slide with isopropanol. Then, rinse them again with deionized water. Dry the PDMS microfluidic channel and the glass slide with compressed air at 1 bar for 1 min. 
	NOTE: The porous structure of the PDMS must be completely dry for the bonding to be effective.</li>
	<li>Place the glass slide and the microfluidic channel in a plasma cleaner (see Table of Materials) and ensure the surfaces to be bonded are facing up. Turn on the plasma cleaner and treat the microfluidic channel and glass slide with air plasma at an airflow of 1 SL/h (standard liter per hour) for 1 min. Bond the microfluidic channel to the glass slide immediately after taking them out of the cleaner by putting them in contact with each other. 
	NOTE: Ensure not to touch the treated surfaces, as this might affect the bonding. When fabricating a microfluidic device with multiple microfluidic channels, expose the microfluidic channels simultaneously and bond them in a single step.</li>
	<li>Place the bonded microfluidic device on an 80 &#176;C hot plate for at least 15 min.</li>
	<li>Store the microfluidic device in a clean cell culture dish until the experiment starts.</li>
</ol>
3. Preparation of the bacterial suspension
<ol>
	<li>Grow a population of Bacillus subtilis NCIB 3610 20 h prior to the start of the experiment by directly inoculating 3 mL of nutrient broth no. 3 culture medium (see Table of Materials) from a frozen glycerol stock in a 15 mL culture tube. Incubate in a shaking incubator at 30 &#176;C and 200 rpm overnight (for 16 h).</li>
	<li>Make a subculture from the overnight culture 4 h prior to the start of the experiment by adding 3 &#181;L of the overnight culture in 3 mL of fresh culture medium (1:1,000 dilution) in a 15 mL culture tube. Incubate the subculture in a shaking incubator at 30 &#176;C at 200 rpm for 3.5-4 h to obtain an optical density at 600 nm (OD600) of 0.1.</li>
</ol>
4. Biofilm growth experiment
<ol>
	<li>Turn on the box incubator of the microscope 3 h before the experiment to ensure a stable temperature of 25 &#176;C. Mount the syringe pump and the pressure sensors (see Table of Materials).</li>
	<li>Connect the inlet and outlet tubing to the microfluidic device. Directly insert a needle (with an outer diameter of 0.6 mm) into the inlet tubing to secure the connection between the tubing and the syringe.</li>
	<li>Place the microfluidic device, 30 mL of deionized water, and 30 mL of culture medium in a vacuum desiccator and degas them for at least 1 h. Then, slowly pull the culture medium and the deionized water into two separate 30 mL syringes. 
	NOTE: This step is crucial to prevent bubble formation in the channel while flushing with the culture medium.</li>
	<li>Mount the microfluidic device on the microscope and place the outlet tubing in a waste container.
	<ol>
		<li>Connect the syringe filled with deionized water to the microfluidic channel through the microfluidic tubing and slowly inject the water until it exits from the pressure sensor outlet. Fill the pressure sensor with water and flush all the bubbles from the tubing connecting the microfluidic channel and the pressure sensor. Close the outlet of the pressure sensor with the screws dedicated to the pressure sensor. 
		NOTE: The described filling procedure ensures that the pressure changes at the microfluidic channels&#39; inlet will be precisely recorded. When running an experiment with multiple microfluidic channels, connect each channel to a separate syringe to ensure equal flow conditions in all channels.</li>
	</ol>
	</li>
	<li>Fill the rest of the microfluidic channel with the deionized water.</li>
	<li>Place a 1.2 &#181;m filter (see Table of Materials) on the culture media syringe. Then, remove the water syringe and carefully connect the culture media syringe to the inlet microfluidic tubing. Mount the syringe on the syringe pump and flush the channel with the culture medium at a flow rate of 2 mL/h for 1 h. 
	NOTE: The filter prevents bacterial cells from entering the syringe during loading. Flushing the microfluidic channel with the culture media will remove the remaining bubbles in the porous structure.</li>
	<li>Set the syringe pump at the desired flow rate (here 1 mL/h) during the experiment and set the pressure reading of the pressure sensors to zero. 
	NOTE: By setting the initial pressure reading to zero, only the pressure difference caused by biofilm development during the experiment will be measured.</li>
	<li>Pipette 1 mL of the bacterial culture at an OD600 of 0.1 in a 1.5 mL centrifuge vial. Load the bacterial culture into the microfluidic channel by placing the outlet tube into the centrifuge vial. After waiting for 5 min to remove any potential air bubbles from the tubes&#39; outlet, withdraw 150 &#181;L of bacterial solution at a flow rate of 1 mL/h, until the microfluidic channel is filled with the bacterial culture.</li>
	<li>Carefully remove the culture media syringe filter and place the outlet into the waste container. Leave the bacterial cells at zero-flow conditions in the microfluidic channel for 3 h to allow their surface attachment in the porous medium. 
	NOTE: Leaving the bacterial cells at zero-flow conditions for 3 h was optimized for the attachment of the bacterial strain used while assuring a well-oxygenated bacterial culture. Other bacterial strains might require more or less time.</li>
	<li>To start the experiment, start the flow by setting the syringe pump to the desired flow rate (here 1 mL/h) and start the pressure reading at 1 Hz.</li>
	<li>Acquire images of the growing biofilm&#160;at the desired time interval, optical configuration, and magnification. 
	&#8203;NOTE: In the present study, images at 4x magnification in the bright-field mode in 18 positions spanning the entire domain of the porous medium were acquired every 6 min for 24 h.</li>
</ol>
5. Image analysis
<ol>
	<li>Reconstruct the entire porous medium from the image sequences recorded by stitching the images from the 18 positions using image analysis software (see Table of Materials) and a stitching algorithm24.</li>
	<li>Save the stitched image sequences as a sequence of singular images. 
	NOTE: If the files are too big, at this point, the images can be rescaled and binned to a reasonable size for further processing.</li>
	<li>Create a mask of the pillars of the porous medium to remove them from the analysis.</li>
	<li>Remove the background of the images by dividing them by their background with the image taken at t = 0 h (Figure 2A) and binarize the images at a threshold suitable to segment the biofilm (Figure 2B).</li>
	<li>Compute the saturation of the biofilm by calculating the area covered by the biofilm (number of pixels attributed to the biofilm) compared to the void area of the porous domain (Figure 3).</li>
</ol>

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The authors declare no conflict of interest.

Microfluidic porous media analogs coupled with pressure sensors provide a suitable tool to study biofilm development in porous media. The versatility in the design of the microfluidic porous medium, specifically the arrangement of the pillars, including diameter, irregular shapes, and pore size, allows the investigation of many geometries. These geometries range from single pores to highly complex, irregularly arranged obstacles mimicking different natural (e.g., soils) and industrial (e.g., membranes and filters) porous...

Biofilms - bacterial colonies embedded in a self-secreted matrix of extra-polymeric substances (EPS) - are ubiquitous in natural porous media, such as soils and aquifers1, and technical and medical applications, like bioremediation2, water filtration3 and medical devices4. The biofilm matrix is comprised of polysaccharides, protein fibers, and extracellular DNA5,6, and strongly depends on the microorganisms, the availability of nutrients, as well as the environmental conditions7. Yet, the functions of the matrix are universal; it forms the scaffold of the biofilm structure, protects the microbial community from mechanical and chemical stresses, and is largely responsible for the biofilms&#39; rheological properties5.
In porous media, the growth of biofilms can clog pores, causing the so-called bioclogging. Biofilm development is controlled by the fluid flow and pore size, defined as the distance separating two pillars, of the porous medium8,9,10. Both the pore size and the fluid flow control the nutrient transport and local shear forces. In turn, the growing biofilm clogs pores, affecting the velocity distribution of the fluid11,12,13, the mass transport, and the hydraulic conductivity of the porous medium14,15. The changes in hydraulic conductivity are reflected through increased pressure in confined systems16,17,18,19. Current microfluidic studies in biofilm development and bioclogging focus on studying the impact of flow velocities in homogeneous geometries16,20 (i.e., with a singular pore size)&#160;or heterogeneous porous media12,21,22. However, to disentangle the effects of flow rates and pore size on biofilm development and the resulting pressure changes in the bioclogged porous medium, a highly controllable and versatile experimental platform allowing the study of different porous media geometries and environmental conditions in parallel is required.
The present study introduces a microfluidic platform that combines pressure measurements with simultaneous imaging of the evolving biofilm within the porous medium. Because of its gas-permeability, bio-compatibility, and flexibility in the channel geometry design, a microfluidic device made of polydimethylsiloxane (PDMS) is a suitable tool for studying biofilm development in porous media. Microfluidics allow&#160;the control of physical and chemical conditions (e.g., fluid flow and nutrient concentration) with high precision to mimic the environment of microbial habitats23. Further, microfluidic devices can easily be imaged with micrometric resolution using an optical microscope and coupled with online measurements (e.g., the local pressure).
In this work, the experiments focus on studying the impact of pore size in a homogeneous porous medium analog under controlled imposed flow conditions. The flow of a culture medium is imposed using a syringe pump, and the pressure difference through the microfluidic channel is measured simultaneously with pressure sensors. Biofilm development is initiated by seeding a planktonic culture of Bacillus subtilis in the microfluidic channel. Regular imaging of the evolving biofilm and image analysis allows one to obtain pore scale resolved information on the surface coverage under various experimental conditions. The correlated information of pressure change and the extent of bioclogging provides crucial input for permeability estimations of bioclogged porous media.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Acrodisc 25 mm Syringe Filter, 1.2 &#181;m Versapor Membrane</td>
			<td>Pall Corporation</td>
			<td>PN4190</td>
			<td>1.2 &#181;m filters</td>
		</tr>
		<tr>
			<td>BD 10 mL Syringe (Luer-Lock)</td>
			<td>BD</td>
			<td>300912</td>
			<td>used to fill the channel with deionised water</td>
		</tr>
		<tr>
			<td>Box Incubator</td>
			<td>Life Imaging Services</td>
			<td></td>
			<td>used to have a stable temperature during the biofilm growth experiment</td>
		</tr>
		<tr>
			<td>Cell density meter CO8000</td>
			<td>WPA biowave</td>
			<td></td>
			<td>OD meter</td>
		</tr>
		<tr>
			<td>Centrifuge vial</td>
			<td>Eppendorf</td>
			<td>30120086</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td>CETONI Base 120</td>
			<td>CETONI GmbH</td>
			<td></td>
			<td>syringe pump</td>
		</tr>
		<tr>
			<td>CorelCAD</td>
			<td>CorelDRAW</td>
			<td></td>
			<td>software used to design the microfluidic channel geometries</td>
		</tr>
		<tr>
			<td>Culture tubes (14 mL, sterile)</td>
			<td>greiner bio-one</td>
			<td></td>
			<td>Culture tubes</td>
		</tr>
		<tr>
			<td>Drying oven, VENTI-Line</td>
			<td>VWR</td>
			<td></td>
			<td>Oven to cure the PDMS</td>
		</tr>
		<tr>
			<td>Handy</td>
			<td>Migros</td>
			<td></td>
			<td>Detergent solution</td>
		</tr>
		<tr>
			<td>Hot plate with temperature control</td>
			<td>VRW</td>
			<td></td>
			<td>to cure the PDMS-glass bonding after plasma treatment</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>FIJI&#160;</td>
			<td></td>
			<td>Image analysis software</td>
		</tr>
		<tr>
			<td>Innova 42 Inc Shaker (New Brunswick)</td>
			<td>Eppendorf</td>
			<td></td>
			<td>Incubator</td>
		</tr>
		<tr>
			<td>Isopropanol (&#62; 99.8%)</td>
			<td>Sigma Aldrich</td>
			<td>67-63-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Masterflex transfer tubing</td>
			<td>Masterflex</td>
			<td>HV-06419-05</td>
			<td>0.020&#39;&#39; ID, 0.06&#39;&#39; OD</td>
		</tr>
		<tr>
			<td>Micro Slides, Plain, 75 x 60 mm</td>
			<td>Corning</td>
			<td>2947-75X50</td>
			<td>Glass slides</td>
		</tr>
		<tr>
			<td>Microfluidic pressure sensor (1 bar)</td>
			<td>Elveflow</td>
			<td></td>
			<td>Pressure sensors</td>
		</tr>
		<tr>
			<td>Miltex Biopsy puncher, diameter 1.5 mm</td>
			<td>Integra</td>
			<td></td>
			<td>Puncher to make the inlet and outlet holes of the microfluidic channel</td>
		</tr>
		<tr>
			<td>mrDev600 developer</td>
			<td>Microresist</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon Eclipse Ti2</td>
			<td>Nikon Instruments</td>
			<td></td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Nutrient broth n&#176;3</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Omnifix Syringe with Luer-Lock</td>
			<td>B.Braun</td>
			<td></td>
			<td>syringes of different volume</td>
		</tr>
		<tr>
			<td>Plasma chamber Zepto</td>
			<td>Diener Electronic</td>
			<td>ZEPTO-1&#160;</td>
			<td>used to plasma bond the PDMS and the glass slide</td>
		</tr>
		<tr>
			<td>Precision wipes (Kimtech Science)</td>
			<td>Kimberly Clark</td>
			<td>KCP-7552</td>
			<td>to dry the glass slide</td>
		</tr>
		<tr>
			<td>Scale</td>
			<td>VWR-CH</td>
			<td>611-2605</td>
			<td>used to weigh the elastomer to crosslinking agent ratio</td>
		</tr>
		<tr>
			<td>Silicon wafer (10 cm)</td>
			<td>Silicon Materials Inc.&#160;</td>
			<td></td>
			<td>N//Phos &#60;100&#62; 1-10 &#937; cm</td>
		</tr>
		<tr>
			<td>Spincoater, Spin module SM150</td>
			<td>Sawatec</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SU8 3050 Photoresist</td>
			<td>Kayakuam</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>S&#252;ss MA6 Mask aligner</td>
			<td>SUSS MicroTec Group</td>
			<td></td>
			<td>used to align the chrome-glass mask</td>
		</tr>
		<tr>
			<td>Sylgard 184</td>
			<td>Dow Corning</td>
			<td></td>
			<td>silicone elastomer kit; curing agent</td>
		</tr>
		<tr>
			<td>Techni Etch Cr01</td>
			<td>Technic</td>
			<td></td>
			<td>Technic</td>
		</tr>
		<tr>
			<td>Tissue culture dish 150</td>
			<td>TPP</td>
			<td>93150</td>
			<td></td>
		</tr>
		<tr>
			<td>Trichloro (1H, 1H, 2H, 2H perfluorooctyl) silane</td>
			<td>Sigma Aldrich</td>
			<td>Sigma Aldrich</td>
			<td>used to silanize the silicane wafer</td>
		</tr>
		<tr>
			<td>Veeco Dektak 6 M</td>
			<td>Veeco</td>
			<td></td>
			<td>Profilometer</td>
		</tr>
	</tbody>
</table>

a microfluidic platform to study bioclogging in porous media

Bacterial biofilms are found in several environmental and industrial porous media, including soils and filtration membranes. Biofilms grow under certain flow conditions and can clog pores, thereby redirecting the local fluid flow. The ability of biofilms to clog pores, the so-called bioclogging, can have a tremendous effect on the local permeability of the porous medium, creating a pressure buildup in the system, and impacting the mass flow through it. To understand the interplay between biofilm growth and fluid flow under different physical conditions (e.g., at different flow velocities and pore sizes), in the present study, a microfluidic platform is developed to visualize biofilm development using a microscope under externally-imposed, controlled physical conditions. The biofilm-induced pressure buildup in the porous medium can be measured simultaneously using pressure sensors and, later, correlated with the surface coverage of the biofilm. The presented platform provides a baseline for a systematic approach to investigate bioclogging caused by biofilms in porous media under flow conditions and can be adapted to studying environmental isolates or multispecies biofilms.

For the present study, a microfluidic device with three parallel microfluidic channels with different pore sizes was used (Figure 1) to study biofilm formation in porous media systematically. The biofilm formation process was visualized using bright-field microscopy. The bacterial cells and the biofilm appeared in the images as darker pixels (Figure 2). In addition, a gradual clogging process was observed; during a 24 h experiment, the initially randomly growing...

Watch this Scientific Journal Video about A Microfluidic Platform to Study Bioclogging in Porous Media at JoVE.com

A Microfluidic Platform to Study Bioclogging in Porous Media

The present protocol describes a microfluidic platform to study biofilm development in quasi-2D porous media by combining high-resolution microscopy imaging with simultaneous pressure difference measurements. The platform quantifies the influence of pore size and fluid flow rates in porous media on bioclogging.

Bacterial biofilms are found in several environmental and industrial porous media, including soils and filtration membranes. Biofilms grow under certain ...

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1.8K Views.  ETH Zurich. The present protocol describes a microfluidic platform to study biofilm development in quasi-2D porous media by combining high-resolution microscopy imaging with simultaneous pressure difference measurements. The platform quantifies the influence of pore size and fluid flow rates in porous media on bioclogging.

Video: A Microfluidic Platform to Study Bioclogging in Porous Media

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 ...

Transient Gene Expression in Tobacco using Gibson Assembly and the Gene Gun

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately using complex regulatory strategies including differential promoters and/or signal sequences. Metabolic engineers and synthetic biologists interested in targeting enzymes to a particular organelle are faced with a challenge: For a protein that is to be localized to more than one organelle, the engineer must clone the same gene multiple times. This work presents a solution to this strategy: harnessing alternative splicing of mRNA. This technology takes advantage of established chloroplast and peroxisome targeting sequences and combines them into a single mRNA that is alternatively spliced. Some splice variants are sent to the chloroplast, some to the peroxisome, and some to the cytosol. Here the system is designed for multiple-organelle targeting with alternative splicing. In this work, GFP was expected to be expressed in the chloroplast, cytosol, and peroxisome by a series of rationally designed 5&#8217; mRNA tags. These tags have the potential to reduce the amount of cloning required when heterologous genes need to be expressed in multiple subcellular organelles. The constructs were designed in previous work11, and were cloned using Gibson assembly, a ligation independent cloning method that does not require restriction enzymes. The resultant plasmids were introduced into Nicotiana benthamiana epidermal leaf cells with a modified Gene Gun protocol. Finally, transformed leaves were observed with confocal microscopy.

This work describes a novel method for selectively targeting subcellular organelles in plants, assayed using the BioRad Gene Gun.

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately ...

A Protocol for Conducting Rainfall Simulation to Study Soil Runoff

Rainfall is a driving force for the transport of environmental contaminants from agricultural soils to surficial water bodies via surface runoff. The objective of this study was to characterize the effects of antecedent soil moisture content on the fate and transport of surface applied commercial urea, a common form of nitrogen (N) fertilizer, following a rainfall event that occurs within 24&#160;hr&#160;after fertilizer application. Although urea is assumed to be readily hydrolyzed to ammonium and therefore not often available for transport, recent studies suggest that urea can be transported from agricultural soils to coastal waters where it is implicated in harmful algal blooms. A rainfall simulator was used to apply a consistent rate of uniform rainfall across packed soil boxes that had been prewetted to different soil moisture contents. By controlling rainfall and soil physical characteristics, the effects of antecedent soil moisture on urea loss were isolated. Wetter soils exhibited shorter time from rainfall initiation to runoff initiation, greater total volume of runoff, higher urea concentrations in runoff, and greater mass loadings of urea in runoff. These results also demonstrate the importance of controlling for antecedent soil moisture content in studies designed to isolate other variables, such as soil physical or chemical characteristics, slope, soil cover, management, or rainfall characteristics. Because rainfall simulators are designed to deliver raindrops of similar size and velocity as natural rainfall, studies conducted under a standardized protocol can yield valuable data that, in turn, can be used to develop models for predicting the fate and transport of pollutants in runoff.

A rainfall simulator was used to apply a consistent rate of uniform rainfall to packed soil boxes in a study of the fate and transport of urea, a nonpoint source environmental contaminant. Under uniform soil and rainfall conditions, antecedent soil moisture content exerted strong control over urea loss in surface runoff.

Rainfall is a driving force for the transport of environmental contaminants from agricultural soils to surficial water bodies via surface runoff. The ...

LC-MS Analysis of Human Platelets as a Platform for Studying Mitochondrial Metabolism

Perturbed mitochondrial metabolism has received renewed interest as playing a causative role in a range of diseases. Probing alterations to metabolic pathways requires a model in which external factors can be well controlled, allowing for reproducible and meaningful results. Many studies employ transformed cellular models for these purposes; however, metabolic reprogramming that occurs in many cancer cell lines may introduce confounding variables. For this reason primary cells are desirable, though attaining adequate biomass for metabolic studies can be challenging. Here we show that human platelets can be utilized as a platform to carry out metabolic studies in combination with liquid chromatography-tandem mass spectrometry analysis. This approach is amenable to relative quantification and isotopic labeling to probe the activity of specific metabolic pathways. Availability of platelets from individual donors or from blood banks makes this model system applicable to clinical studies and feasible to scale up. Here we utilize isolated platelets to confirm previously identified compensatory metabolic shifts in response to the complex I inhibitor rotenone. More specifically, a decrease in glycolysis is accompanied by an increase in fatty acid oxidation to maintain acetyl-CoA levels. Our results show that platelets can be used as an easily accessible and medically relevant model to probe the effects of xenobiotics on cellular metabolism.

Here we show isolated human platelets can be used as an accessible ex vivo model to study metabolic adaptations in response to the complex I inhibitor rotenone. This approach employs isotopic tracing and relative quantification by liquid chromatography-mass spectrometry and can be applied to a variety of study designs.

Perturbed mitochondrial metabolism has received renewed interest as playing a causative role in a range of diseases. Probing alterations to metabolic ...

Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered &#34;bee-safe,&#34; there is mounting evidence that fungicide residues in pollen are associated with bee declines (for both honey and bumble bee species). While the mechanisms remain relatively unknown, researchers have speculated that bee-microbe symbioses are involved. Microbes play a pivotal role in the preservation and/or processing of pollen, which serves as nutrition for larval bees. By altering the microbial community, it is likely that fungicides disrupt these microbe-mediated services, and thereby compromise bee health. This manuscript describes the protocols used to investigate the indirect mechanism(s) by which fungicides may be causing colony decline. Cage experiments exposing bees to fungicide-treated flowers have already provided the first evidence that fungicides cause profound colony losses in a native bumble bee (Bombus impatiens). Using field-relevant doses of fungicides, a series of experiments have been developed to provide a finer description of microbial community dynamics of fungicide-exposed pollen. Shifts in the structural composition of fungal and bacterial assemblages within the pollen microbiome are investigated by next-generation sequencing and metagenomic analysis. Experiments developed herein have been designed to provide a mechanistic understanding of how fungicides affect the microbiome of pollen-provisions. Ultimately, these findings should shed light on the indirect pathway through which fungicides may be causing colony declines.

Microbial consortia within bumble bee hives enrich and preserve pollen for bee larvae. Using next generation sequencing, along with laboratory and field-based experiments, this manuscript describes protocols used to test the hypothesis that fungicide residues alter the pollen microbiome, and colony demographics, ultimately leading to colony loss.

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered ...

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 ...

Microfluidic Devices for Characterizing Pore-scale Event Processes in Porous Media for Oil Recovery Applications

Microfluidic devices are versatile tools for studying transport processes at a microscopic scale. A demand exists for microfluidic devices that are resistant to low molecular-weight oil components, unlike traditional polydimethylsiloxane (PDMS) devices. Here, we demonstrate a facile method for making a device with this property, and we use the product of this protocol for examining the pore-scale mechanisms by which foam recovers crude oil. A pattern is first designed using computer-aided design (CAD) software and printed on a transparency with a high-resolution printer. This pattern is then transferred to a photoresist via a lithography procedure. PDMS is cast on the pattern, cured in an oven, and removed to obtain a mold. A thiol-ene crosslinking polymer, commonly used as an optical adhesive (OA), is then poured onto the mold and cured under UV light. The PDMS mold is peeled away from the optical adhesive cast. A glass substrate is then prepared, and the two halves of the device are bonded together. Optical adhesive-based devices are more robust than traditional PDMS microfluidic devices. The epoxy structure is resistant to swelling by many organic solvents, which opens new possibilities for experiments involving light organic liquids. Additionally, the surface wettability behavior of these devices is more stable than that of PDMS. The construction of optical adhesive microfluidic devices is simple, yet requires incrementally more effort than the making of PDMS-based devices. Also, though optical adhesive devices are stable in organic liquids, they may exhibit reduced bond-strength after a long time. Optical adhesive microfluidic devices can be made in geometries that act as 2-D micromodels for porous media. These devices are applied in the study of oil displacement to improve our understanding of the pore-scale mechanisms involved in enhanced oil recovery and aquifer remediation.

The goal of this procedure is to easily and rapidly produce a microfluidic device with customizable geometry and resistance to swelling by organic fluids for oil recovery studies. A polydimethylsiloxane mold is first generated, and then used to cast the epoxy-based device. A representative displacement study is reported.

Microfluidic devices are versatile tools for studying transport processes at a microscopic scale. A demand exists for microfluidic devices that are ...

Ecosystem Fabrication (EcoFAB) Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A better mechanistic understanding of these complex plant-microbe interactions will be crucial to improving plant production as well as performing basic ecological studies investigating plant-soil-microbe interactions. Here, a detailed description for ecosystem fabrication is presented, using widely available 3D printing technologies, to create controlled laboratory habitats (EcoFABs) for mechanistic studies of plant-microbe interactions within specific environmental conditions. Two sizes of EcoFABs are described that are suited for the investigation of microbial interactions with various plant species, including Arabidopsis thaliana, Brachypodium distachyon, and Panicum virgatum. These flow-through devices allow for controlled manipulation and sampling of root microbiomes, root chemistry as well as imaging of root morphology and microbial localization. This protocol includes the details for maintaining sterile conditions inside EcoFABs and mounting independent LED light systems onto EcoFABs. Detailed methods for addition of different forms of media, including soils, sand, and liquid growth media coupled to the characterization of these systems using imaging and metabolomics are described. Together, these systems enable dynamic and detailed investigation of plant and plant-microbial consortia including the manipulation of microbiome composition (including mutants), the monitoring of plant growth, root morphology, exudate composition, and microbial localization under controlled environmental conditions. We anticipate that these detailed protocols will serve as an important starting point for other researchers, ideally helping create standardized experimental systems for investigating plant-microbe interactions.

Ecosystem Fabrication EcoFAB Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

This article describes detailed protocols for ecosystem fabrication of devices (EcoFABs) that enable the studies of plants and plant-microbe interactions in highly controlled laboratory conditions.

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A ...

Metal Corrosion and the Efficiency of Corrosion Inhibitors in Less Conductive Media

Material corrosion can be a limiting factor for different materials in many applications. Thus, it is necessary to better understand corrosion processes, prevent them and minimize the damages associated with them. One of the most important characteristics of corrosion processes is the corrosion rate. The measurement of corrosion rates is often very difficult or even impossible especially in less conductive, non-aqueous environments such as biofuels. Here, we present five different methods for the determination of corrosion rates and the efficiency of anti-corrosion protection in biofuels: (i) a static test, (ii) a dynamic test, (iii) a static test with a reflux cooler and electrochemical measurements (iv) in a two-electrode arrangement and (v) in a three-electrode arrangement. The static test is advantageous due to its low demands on material and instrumental equipment. The dynamic test allows for the testing of corrosion rates of metallic materials at more severe conditions. The static test with a reflux cooler allows for the testing in environments with higher viscosity (e.g., engine oils) at higher temperatures in the presence of oxidation or an inert atmosphere. The electrochemical measurements provide a more comprehensive view on corrosion processes. The presented cell geometries and arrangements (the two-electrode and three-electrode systems) make it possible to perform measurements in biofuel environments without base electrolytes that could have a negative impact on the results and load them with measurement errors. The presented methods make it possible to study the corrosion aggressiveness of an environment, the corrosion resistance of metallic materials, and the efficiency of corrosion inhibitors with representative and reproducible results. The results obtained using these methods can help to understand corrosion processes in more detail to minimize the damages caused by corrosion.

The testing of processes associated with material corrosion can often be difficult especially in non-aqueous environments. Here, we present different methods for short-term and long-term testing of corrosion behavior of non-aqueous environments such as biofuels, especially those containing bioethanol.

Material corrosion can be a limiting factor for different materials in many applications. Thus, it is necessary to better understand corrosion processes, ...

A Flexible Low Cost Hydroponic System for Assessing Plant Responses to Small Molecules in Sterile Conditions

A wide range of studies in plant biology are performed using hydroponic cultures. In this work, an in vitro hydroponic growth system designed for assessing plant responses to chemicals and other substances of interest is presented. This system is highly efficient in obtaining homogeneous and healthy seedlings of the C3 and C4 model species Arabidopsis thaliana and Setaria viridis, respectively. The sterile cultivation avoids algae and microorganism contamination, which are known limiting factors for plant normal growth and development in hydroponics. In addition, this system is scalable, enabling the harvest of plant material on a large scale with minor mechanical damage, as well as the harvest of individual parts of a plant if desired. A detailed protocol demonstrating that this system has an easy and low-cost assembly, as it uses pipette racks as the main platform for growing plants, is provided. The feasibility of this system was validated using Arabidopsis seedlings to assess the effect of the drug AZD-8055, a chemical inhibitor of the target of rapamycin (TOR) kinase. TOR inhibition was efficiently detected as early as 30 min after an AZD-8055 treatment in roots and shoots. Furthermore, AZD-8055-treated plants displayed the expected starch-excess phenotype. We proposed this hydroponic system as an ideal method for plant researchers aiming to monitor the action of plant inducers or inhibitors, as well as to assess metabolic fluxes using isotope-labeling compounds which, in general, requires the use of expensive reagents.

A simple, versatile, and low-cost in vitro hydroponic system was successfully optimized, enabling large-scale experiments under sterile conditions. This system facilitates the application of chemicals in a solution and their efficient absorption by roots for molecular, biochemical, and physiological studies.