Authors thank Mr. Kotaro Okubo for the kind assistance with SEM imaging. This work was supported by the Japan Society for the Promotion of Science Grant-in-Aid for Basic Research (B) (20310069), Grant-in-Aid for Research Activity Start-up (25880021), and by research grants from the Kurata Memorial Hitachi Science and Technology Foundation and the Nippon Sheet Glass Foundation for Materials Science and Engineering.

1. Preparation of TiO2-coated Glass Coverslip
<ol>
	<li>Number the coverslips using a diamond scriber. This helps not only to keep track of each coverslips but also to ensure that the correct side of the sample is facing up. Clean the coverslips, first under running ddH2O, then by immersing them in piranha solution (H2SO4:H2O2 = 4:1). After 10 min, rinse the coverslips thoroughly, 8 times in ddH2O. Dry the coverslips under N2 flow.</li>
	<li>Set TiO2 target in the radio-frequency (RF) sputtering system. Attach the coverslips onto a sample holder of the sputtering equipment using a polyimide tape. Place the sample holder in the sputtering chamber. Evacuate the chamber until the pressure reaches 2.0&#215;10-4 Pa.</li>
	<li>Introduce Ar gas into the chamber and set the deposition pressure to 4.0 mTorr. While keeping the shutter closed, gradually increase the RF power to 70 W.</li>
	<li>Open the shutter and sputter for 15 min to obtain a film with a thickness of 120-150 nm (Figure 1: Step A). Growth rate of the film needs to be derived for each machine.</li>
</ol>
2. Surface Coating with Cell-repellent Film
<ol>
	<li>Hydrophilize the TiO2 surface by treating the sample with O2 plasma, following instructions provided by the manufacturer of the plasma reactor. We treat the sample for 5 min at 200 W with O2 flow of 100 sccm. Immerse the sample in ddH2O and confirm that the surface is superhydrophilic. Dry the surface thoroughly under N2 flow.</li>
	<li>Prepare 1 mM octadecyltrichlorosilane (OTS) solution by adding 39.6 &#956;l OTS to 100 ml toluene. Immerse the sample in the solution for 1 hr at RT. Conduct this step inside an N2-filled glove bag (Figure 1: Step B).</li>
	<li>To remove physisorbed molecules, sonicate the sample in toluene, acetone, ethanol, and ddH2O for 5 min each, in that order. Rinse the sample four times in fresh ddH2O and dry the surface under N2 flow. The surface should be hydrophobic with a contact angle of 100-110&#176;.</li>
</ol>
3. Ex-Situ Surface Patterning
<ol>
	<li>Working in a laminar flow hood, draw several scratch marks with a diamond scriber on the surface. The marks help in keeping track of the processed regions and also bringing microscopes into focus. Sterilize the OTS-coated TiO2 by immersing the sample in 70% ethanol for 5 min. Then rinse the sample twice in sterilized ddH2O.</li>
	<li>Place the sample in a 35-mm dish, and add 2 ml of PC12 growth medium (see Step 4.2). Incubate for over 3 hr in a CO2 incubator (37 &#176;C). This procedure is intended to let serum albumins absorb onto the surface. Adsorbed albumins inhibit subsequent adsorption of other proteins and cells (Figure 1: Step C).</li>
	<li>While waiting, set up the inverted fluorescence microscope.
	<ol>
		<li>Turn on the arc lamp, insert the UV filter cube, and set the objective lens to 20X.</li>
		<li>Measure light intensity I (W cm-2) using an UV-intensity meter, and calculate irradiation time t (in seconds) for a dose of d (in J cm-2) as: t = d/I. 
		For instance, to irradiate at a dose of 200 J cm-2 using a light source of 600 mW cm-2, 333 sec of irradiation is required.</li>
		<li>Use a stage micrometer and close the field diaphragm to set the size of the region to be irradiated, e.g., 200 &#956;m.</li>
	</ol>
	</li>
	<li>After the 3 hr incubation, supplement the medium with 200 &#956;l of 3.0 mg ml-1 type-IV collagen (Col-IV; final concentration 300 &#956;g ml-1).</li>
	<li>Transfer the 35-mm dish to the microscope stage. Find the scratch mark, focus the microscope onto the sample surface, and irradiate UV-light at a dose of 200 J cm-2 (Figure 1: Step D). The area of UV irradiation can be altered either by adjusting the opening of the field diaphragm or by replacing the field diaphragm with a metal mask of arbitral geometry.</li>
	<li>Replace the medium with a fresh growth medium (without Col-IV), and place the sample back in the incubator.</li>
</ol>
4. Cell Culture
<ol>
	<li>Routine culture of PC12 cells is carried out on a collagen-coated plastic dish.
	<ol>
		<li>To coat a 60-mm tissue-culture dish with Col-IV, first wet the surface with ddH2O and aspirate all ddH2O.</li>
		<li>Prepare 300 &#956;g ml-1 Col-IV by diluting the original solution (3 mg ml-1) 10x with ddH2O.</li>
		<li>Add 200 &#956;l of 300 &#956;g ml-1 Col-IV per 60-mm dish. Let the solution spread through the surface.</li>
		<li>Dry the dish in a laminar flow hood for approximately 1 hr.</li>
		<li>Rinse the surface twice with 4 ml of Dulbecco&#39;s phosphate-buffered saline (D-PBS). When not using the coated dish immediately, rinse the dish once with 4 ml of ddH2O. After aspirating ddH2O, store the dish in the incubator. Try not to keep it stored for more than two weeks.</li>
	</ol>
	</li>
	<li>PC12 cells are grown in a growth medium that consists of: Dulbecco&#39;s modified Eagle&#39;s medium (low glucose) + 10% fetal bovine serum + 5% horse serum + 1% penicillin-streptomycin solution. Incubate the cells in a CO2 incubator (37 &#176;C, 5% CO2), and replace half of the medium every other day. Passage cells before they reach confluence.
	<ol>
		<li>Aspirate the growth medium, and add 2 ml of PBS+ (D-PBS + 10 mg ml-1 bovine serum albumin (BSA) + 10 mM EDTA) prewarmed to 37 &#176;C. Incubate for 5 min at 37 &#176;C. Gently tap the dish to detach all cells.</li>
		<li>Collect cells in a 15 ml conical tube. Rinse the dish with 3 ml of fresh D-PBS, prewarmed to 37 &#176;C.</li>
		<li>Centrifuge the tube at 150 &#215; g for 4 min.</li>
		<li>Aspirate the supernatant, and add 1 ml of the growth medium.</li>
		<li>For routine culture, split the cells 1:3 to 1:5.</li>
	</ol>
	</li>
	<li>To plate cells on the UV-modified OTS/TiO2 sample, count cell density and add 3.0&#215;105 cells in a 35-mm dish (Figure 1: Step E). Incubate the dish in the humidified incubator (37 &#176;C, 5% CO2) for 1-2 days. Both naive and nerve growth factor (NGF)-differentiated PC12 cells can be used for the patterning experiments. For NGF differentiation, 100 ng ml-1 of 7S-NGF is added to the growth medium several days before plating the cells onto the sample. NGF-differentiation seems to increase the adhesibility of the PC12 cells and makes handling easier, especially in the in-situ patterning experiments.</li>
</ol>
5. In-Situ Surface Patterning
<ol>
	<li>After 1-2 days of culture on the ex-situ modified surface, confirm that the cells are attaching and growing only on the UV-irradiated region. Set up the microscope, as described in Step 3.4.</li>
	<li>Transfer the sample to a new 35-mm tissue culture dish containing the growth medium, 100 ng ml-1 NGF (in the case of using NGF-differentiated cells), and 100 &#956;g ml-1 Col-IV.</li>
	<li>Place the dish on the microscope stage. Find the appropriate position, and irradiate UV light at a dose of 200 J cm-2 (Figure 1: Step F, G). The newly created permissive region is indicated with an asterisk in Figure 1: Step G.</li>
	<li>Try to complete processing of a single sample within 30 min. After irradiation, transfer the sample back into the medium without Col-IV.</li>
	<li>Be extremely careful when carrying the dish with cultured cells and transferring the sample from dish to dish to prevent detaching of the patterned cells.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/53045/53045fig1highres.jpg" width="700" /> 
Figure 1. Schematic illustration of the overall process. See text for details. <a href="https://www.jove.com/files/ftp_upload/53045/53045fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have nothing to disclose.

In our current protocol, TiO2 film was formed by RF-magnetron sputtering. We favor this method of deposition since it allows us to reproducibly prepare a photocatalytic TiO2 film with a sub-nm roughness. Although sputter deposition processes are familiar to materials scientists and electronic engineers, it may not be quite accessible to biologists. In that case, spin-coated TiO2 film would be an alternative choice23. In this method, TiO2 nanoparticles dissolved in a ...

Semiconductor lithography processes and its derivatives &#8212; such as photolithography1,2, electron-beam lithography3-6, and microcontact printing7-10 &#8212; have now become an established tool in cell biology to grow living cells in a defined position and geometry. The patterning method relies on the use of microfabricated substrates, consisting of micro-island of cell permissive coating in a non-permissive background. Such substrate serves as a template to pattern the cells. These technologies have provided us the novel methods to engineer cells and their function at a single- and multi-cellular level, to extract the intrinsic properties of cells, and to increase the throughput of cell-based drug screening11.
The degree-of-freedom in cell patterning would greatly increase if the template pattern geometry could be altered in situ, i.e., while cells are cultured on the surface. The conventional methods for pattern formation cannot be directly applied here, since they process samples in atmosphere or in vacuum. Therefore various new surface modification techniques have been proposed, which are based, e.g., on photoreactive compounds12,13 or laser ablation5,14, just to name a few. The proposed methods have been nicely reviewed by Robertus et al.15, and more recently by Choi et al.16 and by Nakanishi17.
Here in this article, we describe a novel protocol of in-situ surface modification, which takes advantage of photocatalytic decomposition of organic molecules on a titanium dioxide (TiO2) surface18,19. In this method, a TiO2 film is inserted between the glass substrate and the organic film that interfaces the cells, and the organic film is decomposed in situ by locally irradiating ultraviolet (UV) light to a region of interest (&#955; &#60; 388 nm). We show that the new protocol can be used to create micropatterns of extracellular matrix proteins and living cells both ex situ and in situ. TiO2 is biocompatible, chemically stable, and optically transparent, features of which makes it friendly to introduce in cell-culture experiments. This protocol provides a materials science-based alternative for modifying cell-culture scaffolds in cell-culture environment.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Glass coverslip</td>
			<td>Warner Instruments</td>
			<td>CS-15R15</td>
			<td>15 mm, #1.5 thickness</td>
		</tr>
		<tr>
			<td>Diamond scriber</td>
			<td>Ogura Jewel Industry</td>
			<td>D-Point Pen</td>
			<td></td>
		</tr>
		<tr>
			<td>RF sputtering system</td>
			<td>ANELVA</td>
			<td>SPC350</td>
			<td></td>
		</tr>
		<tr>
			<td>TiO2 sputtering target</td>
			<td>Kojundo Chemical Lab</td>
			<td>Titanium (IV) oxide, target</td>
			<td>Purity, 99.9%</td>
		</tr>
		<tr>
			<td>Plasma reactor</td>
			<td>Yamato</td>
			<td>PR301</td>
			<td></td>
		</tr>
		<tr>
			<td>n-octadecyltrichlorosilane 
			(OTS)</td>
			<td>Aldrich</td>
			<td>104817</td>
			<td></td>
		</tr>
		<tr>
			<td>Toluene</td>
			<td>Wako</td>
			<td>204-01866</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue-culture dish (35 mm)</td>
			<td>Greiner</td>
			<td>627160</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue-culture dish (60 mm)</td>
			<td>BD Falcon</td>
			<td>353002</td>
			<td></td>
		</tr>
		<tr>
			<td>Type-IV collagen</td>
			<td>Nitta Gelatin</td>
			<td>Cellmatrix Type IV</td>
			<td></td>
		</tr>
		<tr>
			<td>D-PBS</td>
			<td>Gibco</td>
			<td>14190-144</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s modified Eagle&#39;s medium (DMEM)</td>
			<td>Gibco</td>
			<td>11885-084</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum</td>
			<td>Gibco</td>
			<td>12483-020</td>
			<td>Heat-inactivate and pass through a 0.22 mm filter before use</td>
		</tr>
		<tr>
			<td>Horse serum</td>
			<td>Gibco</td>
			<td>26050-088</td>
			<td>Pass through a 0.22 mm filter before use</td>
		</tr>
		<tr>
			<td>Penicillin-streptomycin (100x)</td>
			<td>Nacalai tesque</td>
			<td>26253-84</td>
			<td></td>
		</tr>
		<tr>
			<td>7S nerve growth factor (NGF)</td>
			<td>Alomone Labs</td>
			<td>N-130</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin (BSA)</td>
			<td>Sigma</td>
			<td>A2153</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Dojindo</td>
			<td>N001</td>
			<td>Stock solution in 0.5 M</td>
		</tr>
		<tr>
			<td>TiO2 nanoparticle</td>
			<td>Tayca</td>
			<td>TKD-701</td>
			<td></td>
		</tr>
	</tbody>
</table>

photopatterning proteins and cells in aqueous environment using tio2 photocatalysis

Organic contaminants adsorbed on the surface of titanium dioxide (TiO2) can be decomposed by photocatalysis under ultraviolet (UV) light. Here we describe a novel protocol employing the TiO2 photocatalysis to locally alter cell affinity of the substrate surface. For this experiment, a thin TiO2 film was sputter-coated on a glass coverslip, and the TiO2 surface was subsequently modified with an organosilane monolayer derived from octadecyltrichlorosilane (OTS), which inhibits cell adhesion. The sample was immersed in a cell culture medium, and focused UV light was irradiated to an octagonal region. When a neuronal cell line PC12 cells were plated on the sample, cells adhered only on the UV-irradiated area. We further show that this surface modification can also be performed in situ, i.e., even when cells are growing on the substrate. Proper modification of the surface required an extracellular matrix protein collagen to be present in the medium at the time of UV irradiation. The technique presented here can potentially be employed in patterning multiple cell types for constructing coculture systems or to arbitrarily manipulate cells under culture.

Figure 2A shows a cross-sectional scanning electron microscopy (SEM) image of the sputter-deposited TiO2 film. From the observation, thickness of the film was estimated to be approximately 150 nm. Noticeable here is the flatness of the deposited TiO2 film. Further analysis by atomic force microscopy (AFM) revealed that the root-mean-square (rms) roughness of the surface was 0.2 nm (Figure 2B).
When the TiO2 surface is modified ...

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Photopatterning Proteins and Cells in Aqueous Environment Using TiO2 Photocatalysis

We describe a protocol for modifying cell affinity of a scaffold surface in aqueous environment. The method takes advantage of titanium dioxide photocatalysis to decompose organic film in the photo-irradiated region. We show that it can be used to create microdomains of scaffolding proteins, both ex situ and in situ.

Photopatterning Proteins and Cells in Aqueous Environment Using TiO2 Photocatalysis

Organic contaminants adsorbed on the surface of titanium dioxide (TiO2) can be decomposed by photocatalysis under ultraviolet (UV) light. Here we describe ...

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Research

JoVE Journal

Bioengineering

7.8K Views.  Tohoku University. We describe a protocol for modifying cell affinity of a scaffold surface in aqueous environment. The method takes advantage of titanium dioxide photocatalysis to decompose organic film in the photo-irradiated region. We show that it can be used to create microdomains of scaffolding proteins, both ex situ and in situ.

Video: Photopatterning Proteins and Cells in Aqueous Environment Using TiO2 Photocatalysis

Fabrication of Electrochemical-DNA Biosensors for the Reagentless Detection of Nucleic Acids, Proteins and Small Molecules

As medicine is currently practiced, doctors send specimens to a central laboratory for testing and thus must wait hours or days to 
receive the results. Many patients would be better served by rapid, bedside tests. To this end our laboratory and others have developed a versatile, reagentless 
biosensor platform that supports the quantitative, reagentless, electrochemical detection of nucleic acids (DNA, RNA), proteins (including antibodies) and small 
molecules analytes directly in unprocessed clinical and environmental samples. In this video, we demonstrate the preparation and use of several biosensors in this 
"E-DNA" class. In particular, we fabricate and demonstrate sensors for the detection of a target DNA sequence in a polymerase chain reaction mixture, an HIV-specific antibody and the drug cocaine. The preparation procedure requires only three hours of hands-on effort followed by an overnight incubation, and their use requires only minutes.

"E-DNA" sensors, reagentless, electrochemical biosensors that perform well even when challenged directly in blood and other complex matrices, have been adapted to the detection of a wide range of nucleic acid, protein and small molecule analytes. Here we present a general procedure for the fabrication and use of such sensors.

As medicine is currently practiced, doctors send specimens to a central laboratory for testing and thus must wait hours or days to 
receive the results. ...

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural biology 1. In recent years, this method has achieved great success in studying structures of proteins and macromolecular complexes 2, 3. Compared with electron cryomicroscopy (cryoEM), in which frozen hydrated protein samples are embedded in a thin layer of vitreous ice 4, negative staining is a simpler sample preparation method in which protein samples are embedded in a thin layer of dried heavy metal salt to increase specimen contrast 5. The enhanced contrast of negative stain EM allows examination of relatively small biological samples. In addition to determining three-dimensional (3D) structure of purified proteins or protein complexes 6, this method can be used for much broader purposes. For example, negative stain EM can be easily used to visualize purified protein samples, obtaining information such as homogeneity/heterogeneity of the sample, formation of protein complexes or large assemblies, or simply to evaluate the quality of a protein preparation.
In this video article, we present a complete protocol for using an EM to observe negatively stained protein sample, from preparing carbon coated grids for negative stain EM to acquiring images of negatively stained sample in an electron microscope operated at 120kV accelerating voltage. These protocols have been used in our laboratory routinely and can be easily followed by novice users.

Visualizing protein samples by negative stain electron microscopy (EM) has become a popular structural analysis method. It is useful for quantitative structural analysis, such as calculating a 3D reconstruction of the molecules being studied, and also for qualitative examination of the quality of protein preparations. In this article we present detailed protocols for preparing the EM grids, staining the sample and visualizing the sample in an electron microscope. Novice users can follow these protocols easily and to utilize negative stain EM as a routine assay, in addition to other biochemical assays, for evaluating their protein samples.

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural ...

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

A Toolkit to Enable Hydrocarbon Conversion in Aqueous Environments

This work puts forward a toolkit that enables the conversion of alkanes by Escherichia coli and presents a proof of principle of its applicability. The toolkit consists of multiple standard interchangeable parts (BioBricks)9 addressing the conversion of alkanes, regulation of gene expression and survival in toxic hydrocarbon-rich environments.
A three-step pathway for alkane degradation was implemented in E. coli to enable the conversion of medium- and long-chain alkanes to their respective alkanols, alkanals and ultimately alkanoic-acids. The latter were metabolized via the native &beta;-oxidation pathway. To facilitate the oxidation of medium-chain alkanes (C5-C13) and cycloalkanes (C5-C8), four genes (alkB2, rubA3, rubA4and rubB) of the alkane hydroxylase system from Gordonia sp. TF68,21 were transformed into E. coli. For the conversion of long-chain alkanes (C15-C36), theladA gene from Geobacillus thermodenitrificans was implemented. For the required further steps of the degradation process, ADH and ALDH (originating from G. thermodenitrificans) were introduced10,11. The activity was measured by resting cell assays. For each oxidative step, enzyme activity was observed.
To optimize the process efficiency, the expression was only induced under low glucose conditions: a substrate-regulated promoter, pCaiF, was used. pCaiF is present in E. coli K12 and regulates the expression of the genes involved in the degradation of non-glucose carbon sources.
The last part of the toolkit - targeting survival - was implemented using solvent tolerance genes, PhPFD&alpha; and &beta;, both from Pyrococcus horikoshii OT3. Organic solvents can induce cell stress and decreased survivability by negatively affecting protein folding. As chaperones, PhPFD&alpha; and &beta; improve the protein folding process e.g. under the presence of alkanes. The expression of these genes led to an improved hydrocarbon tolerance shown by an increased growth rate (up to 50%) in the presences of 10% n-hexane in the culture medium were observed.
Summarizing, the results indicate that the toolkit enables E. coli to convert and tolerate hydrocarbons in aqueous environments. As such, it represents an initial step towards a sustainable solution for oil-remediation using a synthetic biology approach.

A sustainable auto regulating bacterial system for the remediation of oil pollutions was designed using standard interchangeable DNA parts (BioBricks). An engineered E. coli strain was used to degrade alkanes via &beta;-oxidation in toxic aqueous environments. The respective enzymes from different species showed alkane degradation activity. Additionally, an increased tolerance to n-hexane was achieved by introducing genes from alkane-tolerant bacteria.

This work puts forward a toolkit that enables the conversion of alkanes by Escherichia coli and presents a proof of principle of its applicability. The ...

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, phosphorus, sulfate and trace elements that plants acquire from the soil. If we want to develop sustainable approaches to producing high crop yield, we need to better understand how the root develops, takes up a wide spectrum of nutrients, and interacts with symbiotic and pathogenic organisms. To accomplish these goals, we need to be able to explore roots in microscopic detail over time periods ranging from minutes to days.
We developed the RootChip, a polydimethylsiloxane (PDMS)- based microfluidic device, which allows us to grow and image roots from Arabidopsis seedlings while avoiding any physical stress to roots during preparation for imaging1 (Figure 1). The device contains a bifurcated channel structure featuring micromechanical valves to guide the fluid flow from solution inlets to each of the eight observation chambers2. This perfusion system allows the root microenvironment to be controlled and modified with precision and speed. The volume of the chambers is approximately 400 nl, thus requiring only minimal amounts of test solution.
Here we provide a detailed protocol for studying root biology on the RootChip using imaging-based approaches with real time resolution. Roots can be analyzed over several days using time lapse microscopy. Roots can be perfused with nutrient solutions or inhibitors, and up to eight seedlings can be analyzed in parallel. This system has the potential for a wide range of applications, including analysis of root growth in the presence or absence of chemicals, fluorescence-based analysis of gene expression, and the analysis of biosensors, e.g. FRET nanosensors3.

This article provides a protocol for cultivation of Arabidopsis seedlings in the RootChip, a microfluidic imaging platform that combines automated control of growth conditions with microscopic root monitoring and FRET-based measurement of intracellular metabolite levels.

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, ...

Cell Co-culture Patterning Using Aqueous Two-phase Systems

Cell patterning technologies that are fast, easy to use and affordable will be required for the future development of high throughput cell assays, platforms for studying cell-cell interactions and tissue engineered systems. This detailed protocol describes a method for generating co-cultures of cells using biocompatible solutions of dextran (DEX) and polyethylene glycol (PEG) that phase-separate when combined above threshold concentrations. Cells can be patterned in a variety of configurations using this method. Cell exclusion patterning can be performed by printing droplets of DEX on a substrate and covering them with a solution of PEG containing cells. The interfacial tension formed between the two polymer solutions causes cells to fall around the outside of the DEX droplet and form a circular clearing that can be used for migration assays. Cell islands can be patterned by dispensing a cell-rich DEX phase into a PEG solution or by covering the DEX droplet with a solution of PEG. Co-cultures can be formed directly by combining cell exclusion with DEX island patterning. These methods are compatible with a variety of liquid handling approaches, including manual micropipetting, and can be used with virtually any adherent cell type.

Aqueous two-phase systems were used to simultaneously pattern multiple populations of cells. This fast and easy method for cell patterning takes advantage of the phase separation of aqueous solutions of dextran and polyethylene glycol and the interfacial tension that exists between the two polymer solutions.

Cell patterning  technologies that are fast, easy to use and affordable will be required for the  future development of high throughput cell assays, ...

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range of 10-100 &#197;. By measuring the distances under various ligated conditions, conformational changes of the protein can be easily assessed. With LRET, a lanthanide, most often chelated terbium, is used as the donor fluorophore, affording advantages such as a longer donor-only emission lifetime, the flexibility to use multiple acceptor fluorophores, and the opportunity to detect sensitized acceptor emission as an easy way to measure energy transfer without the risk of also detecting donor-only signal. Here, we describe a method to use LRET on membrane proteins expressed and assayed on the surface of intact mammalian cells. We introduce a protease cleavage site between the LRET fluorophore pair. After obtaining the original LRET signal, cleavage at that site removes the specific LRET signal from the protein of interest allowing us to quantitatively subtract the background signal that remains after cleavage. This method allows for more physiologically relevant measurements to be made without the need for purification of protein.

We describe here an improved Luminescence Resonance Energy Transfer (LRET) method where we introduce a protease cleavage site between the donor and acceptor fluorophore sites. This modification allows us to obtain specific LRET signals arising from membrane proteins of interest, allowing for the study of membrane proteins without protein purification.

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range ...

Aqueous Droplets Used as Enzymatic Microreactors and Their Electromagnetic Actuation

For the successful implementation of microfluidic reaction systems, such as PCR and electrophoresis, the movement of small liquid volumes is essential. In conventional lab-on-a-chip-platforms, solvents and samples are passed through defined microfluidic channels with complex flow control installations. The droplet actuation platform presented here is a promising alternative. With it, it is possible to move a liquid drop (microreactor) on a planar surface of a reaction platform (lab-in-a-drop). The actuation of microreactors on the hydrophobic surface of the platform is based on the use of magnetic forces acting on the outer shell of the liquid drops which is made of a thin layer of superhydrophobic magnetite particles. The hydrophobic surface of the platform is needed to avoid any contact between the liquid core and the surface to allow a smooth movement of the microreactor. On the platform, one or more microreactors with volumes of 10 &#181;L can be positioned and moved simultaneously. The platform itself consists of a 3 x 3 matrix of electrical double coils which accommodate either neodymium or iron cores. The magnetic field gradients are automatically controlled. By variation of the magnetic field gradients, the microreactors' magnetic hydrophobic shell can be manipulated automatically to move the microreactor or open the shell reversibly. Reactions of substrates and corresponding enzymes can be initiated by merging the microreactors or bringing them into contact with surface immobilized catalysts.

Lab-in-a-drop reaction systems allow the versatile implementation of complex reactions in a microfluidic scale. An automated actuation platform consisting of a 3 x 3 matrix of electromagnetic coils was developed and successfully used to merge two 10 &#181;L microreactors and thereby initiate an enzymatic reaction in the resulting liquid marbles.

For the successful implementation of microfluidic reaction systems, such as PCR and electrophoresis, the movement of small liquid volumes is essential. In ...

A Versatile Method of Patterning Proteins and Cells

Substrate and cell patterning techniques are widely used in cell biology to study cell-to-cell and cell-to-substrate interactions. Conventional patterning techniques work well only with simple shapes, small areas and selected bio-materials. This article describes a method to distribute cell suspensions as well as substrate solutions into complex, long, closed (dead-end) polydimethylsiloxane (PDMS) microchannels using negative pressure. This method enables researchers to pattern multiple substrates including fibronectin, collagen, antibodies (Sal-1), poly-D-lysine (PDL), and laminin. Patterning of substrates allows one to indirectly pattern a variety of cells. We have tested C2C12 myoblasts, the PC12 neuronal cell line, embryonic rat cortical neurons, and amphibian retinal neurons. In addition, we demonstrate that this technique can directly pattern fibroblasts in microfluidic channels via brief application of a low vacuum on cell suspensions. The low vacuum does not significantly decrease cell viability as shown by cell viability assays. Modifications are discussed for application of the method to different cell and substrate types. This technique allows researchers to pattern cells and proteins in specific patterns without the need for exotic materials or equipment and can be done in any laboratory with a vacuum.

This report describes a simple, easy to perform technique, using low pressure vacuum, to fill microfluidic channels with cells and substrates for biological research.

Substrate and cell patterning techniques are widely used in cell biology to study cell-to-cell and cell-to-substrate interactions. Conventional patterning ...