The authors acknowledge funding from the Australian Research Council and National Health and Medical Research Council of Australia and also like to thank the Australian National Fabrication Facility for the SU-8 master for the microcontact printing. SHN is supported by the Ministry of Higher Education Malaysia and Universiti Sains Malaysia.

1. Passivation of Glass Coverslips with PLL-PEG-biotin
This step is designed to passivate the surface so that cells adhere and migrate onto specific regions on the glass coverslip that are created with microcontract printing (Step 2-3a) or dip pen lithography (Step 3b). For the passivation, a co-polymer of poly-L-lysine (PLL) and polyethylene glycol (PEG) is used in which 20% of PEG molecules are grafted to biotin (PLL-PEG-biotin).
<ol>
 <li>To clean glass coverslips (18 mm x 18 mm), submerge them in a rack in 100% ethanol and place the rack in a sonicating bath for 30 min. Dry the glass coverslips in air or under a stream of nitrogen.</li>
 <li>Next, sonicate the glass coverslips for 30 min in 1 M NaOH. Afterwards rinse the glass surfaces by carefully dipping the rack in a beaker filled with 1.5 L Milli-Q H2O. Repeat the rinsing process three times using fresh Milli-Q H2O each time. Dry the glass coverslips in an oven at 60 &deg;C.</li>
 <li>Place half of the clean glass coverslips individually in a humid chamber, which is made of a 24-well plate partially filled with H2O. Drop 15 &mu;l of PLL-PEG-biotin (1 mg/ml) in PBS on each glass coverslip. Take the other half of the remaining clean glass coverslips and carefully sandwich the PEG solution. Leave the coverslips for 1 hr in the humid chamber. Slide the sandwiched coverslips gently away from each other without scraping the PEG-coated surface and rinse them with Milli-Q H2O. The water should slide off easily on the PEG-treated side. If necessary, air-dry the glass coverslips on paper towel with the treated surface facing upward. Store the glass coverslips in a vacuum desiccator until further use.</li>
</ol>
2. Fabrication of Stamps for Microcontact Printing
We describe two processes to pattern streptavidin tracks onto the passivated glass coverslips. The first one, microcontact printing, typically requires the fabrication of a silicon master from which a PDMS stamp is cast. In our case, the pattern on the PDMS stamp are 100 &mu;m-wide lines that are spaced 290 &mu;m center-to-center with a feature height of 25 &mu;m. Below, we describe briefly how to fabricate a suitable silicon master (Step 2.1-2.3) and PDMS stamp (Step 2.4). However, there are multiple protocols for microcontact printing described in the literature19-23, which are suitable for printing protein patterns onto glass coverslips. Masters can also be custom made and purchased from commercial sources.
<ol>
 <li>First, clean the silicon wafer (4" in diameter) by incubating it in piranha solution (3:1 v:v of sulphuric acid and 30% hydrogen peroxide) for 20 min. Rinse the wafer thoroughly 10 times with MilliQ H2O. Dry the wafer in an oven at 150 &deg;C O/N.</li>
 <li>Rinse the clean and dried silicon wafer with 100% acetone, followed by 100% isopropyl alcohol. Next spin-coat 2 ml of GM1070 SU-8 photoresist (Gersteltec Sarl) onto the wafer at 3,100 rpm for 40 sec to create a 25 &mu;m thick layer. The sample is then baked on a hot plate at 65 &deg;C for 5 min followed by a further 5 min at 95 &deg;C.</li>
 <li>Expose the SU-8 photoresist to UV light for 60 sec under a photomask with the desired pattern (for example using a Quintel Q6000 mask aligner) and repeat the baking process describe in 2.2. Then the sample is developed by immersing the wafer in SU-8 developer (Gersteltec Sarl) for 4 min, rinsed in 100% isopropyl alcohol and dried under nitrogen flow. The area of the SU-8 photoresist that was not exposed to UV light is thus removed, creating the inverse pattern of the PDMS stamp.</li>
 <li>The next step is to cast a PDMS stamp from the silicon master. First, thoroughly mix 1 part (w/w) of the curing agent with 10 parts of elastomer prepolymer (Sylgard 184). To remove any air bubbles, degas the mixture in a desiccator attached to a vacuum line for 1 hr. Next, pour the PDMS elastomer onto the silicon master inside a Petri dish to about 1 cm in height and cure the master and PDMS mixture in an oven at 70 &deg;C for 1 hr. Finally, the PDMS stamp is removed from the silicon master and cut to size.</li>
</ol>
3. Patterning Streptavidin or Adhesive Cues onto Passivated Glass Coverslips
After passivating glass coverslips, protein patterns are printed with microcontact printing (Step 3a) or dip-pen lithography (Step 3b). In our case, streptavidin is printed as lines, which form the basis for RGD gradients (Step 4). The same procedure can also be used for printing matrix proteins to create adhesive patterns.
An alternative method to microcontact printing for the patterning of biomolecules onto surfaces is dip pen lithography. In dip pen lithography, a cantilever is used to transfer a small volume of solution onto the surface. The size of the cantilevers, viscosity of the solution, contact time of the cantilever on the surface, hydrophobicity of the surface and humidity in the printing chamber determine the size of the printed features. The advantage of this method is that different patterns can be printed for each experiment, as there is no need to fabricate masters and stamps. The disadvantage is that it takes longer to print a given pattern and that dip pen lithography requires a specialist instrument that may not be available in many laboratories. The dip pen lithography instrument that we used was the NanoeNabler from Bioforce Nanoscience. Here, we printed small dots that were &lt;10 &mu;m in diameter, for which we used SPT10 cantilevers.
3a. Microcontact Printing (&mu;CP) Technique
<ol>
 <li>To clean and make the PDMS stamp more hydrophilic, treat the patterned side of the PDMS stamps in an UV and ozone (for example, UV Ozone Procleaner from Bioforce Nanoscience) for 1.5 hr. Alternatively, use a plasma cleaner for a shorter time for the same purpose. This process creates an oxidized PDMS surface24, which improves the printing process.</li>
 <li>Immediately after the ozone or plasma treatment, place and spread 10 &mu;l streptavidin (1 mg/ml in PBS) onto the PDMS stamps and leave for 1 hr in a humid chamber (such as a partially filled 6-well plate, see Step 1.3). If tracks should be visible under the fluorescent microscope, use streptavidin that is conjugated to a fluorophore such as streptavidin-AlexaFluor350.</li>
 <li>Remove excess streptavidin from the stamp with tissue paper and air dry the stamp for approximately 1 min. Lightly press the stamp onto the PLL-PEG-biotin-coated glass coverslip.</li>
 <li>If fluorescent streptavidin was used in Step 3.a2, examine the pattern using under an epifluorescence microscope. Store printed surfaces in a desiccator.</li>
</ol>
3b. Dip Pen Lithography
<ol>
 <li>First, mix 1 part of 1 mg/ml streptavidin or streptavidin-AlexaFluor350 with 10 parts of glycerol (v/v) and load 5 &mu;l of the mixture onto the cantilever. Then begin the printing process as described in the operation manual of the dip pen lithography instrument. To reduce the spot size to approximately 5 &mu;m, adjust printing speed, contact time of the cantilever on the surface or vertical distance of the cantilever to the surface. In the example shown in Figure 2c, dots were printed in a line with row and column separation set to 10-15 &mu;m. This distance is sufficient to prevent dots from merging into each other but allows the viewing of multiple dots in a single field of view with most microscope settings.</li>
 <li>Store printed surfaces in a desiccator.</li>
</ol>
4. Creating Adhesive Gradients onto Streptavidin-pattern Surfaces
We create adhesive gradients of biotin-RGD onto glass coverslips that are coated with streptavidin (Figure 3) or contain streptavidin patterns on passivated glass coverslips (Figure 5). To create a surface gradient, we used a commercially available microfluidic device (called sticky-Slide Chemotaxis 3D from Ibidi) that creates a stable solution gradient by passive diffusion. To compete for streptavidin binding sites, we employed two opposing gradients of biotin-RGD and biotin that was not conjugated to RGD (see Figure 1).
<ol>
 <li>Adhere the streptavidin-coated or -patterned glass coverslips onto the sticky side of the sticky-Slide Chemotaxis 3D device. To ensure that there is no leakage for several hours, the edges of the device are sealed with a thin layer of warmed Vaseline and with a second layer of a mixture of 1 part of Vaseline to 1 part of paraffin wax (w/w).</li>
 <li>Fill the channel of the device with 6 &mu;l PBS in the channel. Then fill the two reservoirs with 70 &mu;l of 0.03 &mu;g/&mu;l biotin-4-fluorescein in PBS and 70 &mu;l of 0.03 &mu;g/&mu;l biotin-RGD in PBS, respectively. Incubate the samples at RT in the dark for 1 hr.</li>
 <li>Remove the biotin solutions and carefully rinse the surface while still attached to the microfluidic device twice with PBS. Mark the direction of the adhesive gradient and if needed, store the device filled with PBS at 4 &deg;C for but ensure that the device does not dry out. If required, analyze the adhesive gradient with a fluorescent microscope.</li>
</ol>
5. Labelling of Cells with Fluorophores for Live Cell Imaging
Cells are labelled with fluorescent dyes that assist cell tracking under the fluorescent microscope. Common probes are fluorescent organelle markers, such Syto 64 Red, which labels the cell nucleus. To do so, cells are first washed three times with PBS, then incubated with 1 &mu;M Syto 64 Red in culture media for 45 min and finally washed a further three times with PBS. Please note that cells should not be kept in PBS for long periods of time.
Alternative dyes for cell tracking are CellTracker series that stain the cytoplasm and are retained during cell migration and division. Transfection of cells with fluorescent fusion protein is particularly useful to record the subcellular distributions of a protein of interest during migration. However, fluorescent proteins typically photo-bleach faster than organic dyes so that long-term observations may not be possible.
6. Loading of Cells with a Soluble Gradient into the Microfluidics Device
<ol>
 <li>Count fluorescently labelled cells and split into two tubes of equal cell numbers. Wash and resuspend one tube of cells with media containing the chemoattractant, such as low glucose Dulbecco's Modified Eagle Medium (DMEM) with 10% fetal bovine serum (FBS). Wash and resuspend the other tube of cells in media without the chemoattractant, i.e. DMEM with 0% FBS. The final concentration of cells in each tube should be 5 x 105 cells/ml.</li>
 <li>Remove all the solutions from the microfluidic device (from Step 4.3) and place it on the microscope stage (see Step 7). Load 70 &mu;l of cells (5 x 105 cells/ml in 0% FBS DMEM) into one reservoir and 70 &mu;l of HeLa cells (5 x 105 cells/ml in 10% FBS DMEM) into another. Loosely place a tape over the reservoirs to avoid evaporation. If cells on the surface are pre-incubated prior to imaging, it is advisable that the cells are loaded into the device in growth media without the gradient. The device is then placed on the microscope stage and the media replaced with gradient media i.e. DMEM without FBS in one reservoir and DMEM with 10% FBS in the other.</li>
</ol>
7. Live Cell Imaging and Data Analysis
<ol>
 <li>Switch on the microscope (Nikon Ti-E) and warm up the stage at least 1 hr before starting the experiment.</li>
 <li>Ensure that the channel of the microfluidic device is in the field of view and cells are in focus. Select image acquisition parameter such as exposure times and fluorescent filters, image sequence of bright field and fluorescence that captures cells and tracks as well as time points and recording length (e.g. every 15 min for 16 hr).</li>
 <li>Analyze data with appropriate softwares such as ImageJ Manual Tracking or Ibidi Chemotaxis and Migration Tool.</li>
</ol>

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We have no commercial interests to disclose.

In this protocol, we used a commercially available microfluidic device (sticky-Slide Chemotaxis 3D from Ibidi) to study the effects of chemically modified surfaces and chemoattractant gradients on cell migration. This microfluidic setup does not require flow because the gradient is established by diffusion along the length of the channel. This is important because flow could differentially affect slowly migrating cells such as fibroblasts that form stable focal adhesions and fast migrating cells like most leuk...

Directed cell migration is a fundamental property of many cells and is a key aspect of many normal physiological processes, including embryonic development, defense against infection and wound healing. In addition, cell migration also plays a prominent role in many diseases such as vascular disease, tumour cell metastasis and chronic inflammation1,2. While the classical states of cell migration - polarization, protrusion extension, formation of adhesion, force generation and rear retraction - are generally accepted3,4 , the elucidation of spatiotemporal mechanisms by which signal integration is coordinated has been more challenging.
The extracellular matrix (ECM) serves as a substrate for cell adhesion, and through inherent chemical5 and physical6 diversity, provides directional cues for cell navigation7,8. In addition to these adhesive cues9, soluble factors10-12 such as chemokines and growth factors can induce directed cell migration through chemoattractant receptors and their downstream motogenic signalling. Currently, it is not known whether engagement and signalling through adhesion receptors (i.e. integrins) or chemoattractant receptors, such as receptor tyrosine kinase (RTK), are dominant. Nor is it known whether hierarchies of receptor systems are cell type specific.
Live cell microscopy gives a wealth of information that is not accessible in bulk assays and can be combined with microfluidic devices to generate gradients of immobilized 13,14 and soluble cues15 16. The method described here utilizes a series of simple and established steps for surface modification in conjunction with a commercially available microfluidic chamber to create an imaging assay for cell migration that can easily be implemented in a cell biology laboratory (Figure 1). The assembled microfluidic device has optical qualities that are comparable to glass17 and the gradients of diffusible molecules are stable for at least 16 hr. The system can be used for epifluorescence and advanced microscopy techniques. Unlike other chemotaxis setups18, this system is suitable for recording slowly migrating, adherent cells. Importantly, the system is modular and easily allows the introduction of alternative adhesive or soluble migration cues and examination of various cell types.

<table border="1">
	<tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Coverglass staining outfits</td>
 <td>Thomas Scientific</td>
 <td>8542 E40</td>
 <td>Coverslip rack</td>
 </tr>
 <tr>
 <td>Oven</td>
 <td>Binder</td>
 <td></td>
 <td>ED 53 series</td>
 </tr>
 <tr>
 <td>Silicon wafer</td>
 <td>Silicon Quest</td>
 <td>708-007</td>
 <td>Boron doped &lt;100&gt; wafer, 4" diameter, 500 &mu;m, single side polished</td>
 </tr>
 <tr>
 <td>GM1070 SU-8 photoresist</td>
 <td>Gersteltec Sarl</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>SU-8 developer</td>
 <td>Gersteltec Sarl</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Sylgard 184 curing agent</td>
 <td>Dow Corning</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Sylgard 184 elastomer prepolymer</td>
 <td>Dow Corning</td>
 <td></td>
 <td >&nbsp;</td>
 </tr>
 <tr>
 <td>PLL-PEG-biotin (20%)</td>
 <td>SuSos AG</td>
 <td>PLL(20)-g[3.5]-PEG(2)/PEG(3.4)-Biotin (20%)</td>
 <td>1 mg/ml in PBS</td>
 </tr>
 <tr>
 <td>Fluorescein</td>
 <td>Sigma</td>
 <td>46955</td>
 <td>1 mM in PBS</td>
 </tr>
 <tr>
 <td>Streptavidin-AlexaFluor350</td>
 <td>Invitrogen</td>
 <td>S-11249</td>
 <td>1 mg/ml in PBS</td>
 </tr>
 <tr>
 <td>Biotin-4-fluorescein</td>
 <td>Invitrogen</td>
 <td>B-1370</td>
 <td>0.03 &mu;g/&mu;l in PBS</td>
 </tr>
 <tr>
 <td>Biotin-RGD</td>
 <td>GenScript</td>
 <td>SC1208</td>
 <td>0.03 mg/ml in PBS</td>
 </tr>
 <tr>
 <td>Syto 64 Red</td>
 <td>Invitrogen</td>
 <td> S-11346 </td>
 <td>1 &mu;M in PBS</td>
 </tr>
 <tr>
 <td>Sticky slide chemotaxis 3D</td>
 <td>Ibidi</td>
 <td>80328</td>
 <td></td>
 </tr>
 <tr>
 <td>200 &mu;l Greiner yellow bevelled tip</td>
 <td>Greiner Bio-One</td>
 <td>739261</td>
 <td></td>
 </tr>
 <tr>
 <td>Vaseline</td>
 <td>Sigma</td>
 <td>16415</td>
 <td></td>
 </tr>
 <tr>
 <td>Paraffin wax, mp 55-57 &deg;C</td>
 <td>Sigma</td>
 <td>327204</td>
 <td></td>
 </tr>
 <tr>
 <td>Nano eNabler 10 &mu;m cantilever </td>
 <td>BioForce</td>
 <td>SPT-S-C-10s</td>
 <td></td>
 </tr>
 <tr>
 <td>Image J software</td>
 <td>National Institute of Health</td>
 <td></td>
 <td ><a href="http://rsbweb.nih.gov/ij/download.html" target="_blank">rsbweb.nih.gov/ij/download.html</a></td>
 </tr>
 <tr>
 <td>Manual Tracking plugin</td>
 <td>Fabrice Cordeli&egrave;res</td>
 <td></td>
 <td ><a href="http://rsb.info.nih.gov/ij/plugins/track/track.html" target="_blank">rsb.info.nih.gov/ij/plugins/track/track.html</a></td>
 </tr>
 <tr>
 <td>Chemotaxis and Migration Tool</td>
 <td>Ibidi GmbH</td>
 <td></td>
 <td><a href="http://www.ibidi.com/applications/ap_chemotaxis.html" target="_blank">www.ibidi.com/applications/ap_chemotaxis.html</a></td>
 </tr>
	 </tbody>
 </table>

creating adhesive and soluble gradients for imaging cell migration with fluorescence microscopy

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as growth factors. Here, we outline a method to create opposing gradients of adhesive and soluble cues in a microfluidic chamber, which is compatible with live cell imaging. A copolymer of poly-L-lysine and polyethylene glycol (PLL-PEG) is employed to passivate glass coverslips and prevent non-specific adsorption of biomolecules and cells. Next, microcontact printing or dip pen lithography are used to create tracks of streptavidin on the passivated surfaces to serve as anchoring points for the biotinylated peptide arginine-glycine-aspartic acid (RGD) as the adhesive cue. A microfluidic device is placed onto the modified surface and used to create the gradient of adhesive cues (100% RGD to 0% RGD) on the streptavidin tracks. Finally, the same microfluidic device is used to create a gradient of a chemoattractant such as fetal bovine serum (FBS), as the soluble cue in the opposite direction of the gradient of adhesive cues.

To understand how cell integrate migratory signals25, we have developed a method to image cells with fluorescence microscopy that migrate in an environment with competing adhesive and soluble gradients (Figure 1). Adhesive tracks that contained fluorescent streptavidin and biotinylated RGD were created with microcontact printed tracks and dip pen lithography (Figure 2). Successful microcontact printing is indicated by the line profile of the fluorescence intensity across the t...

Watch this Scientific Journal Video about Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy at JoVE.com

Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy

A method for the assembly of adhesive and soluble gradients in a microscopy chamber for live cell migration studies is described. The engineered environment combines antifouling surfaces and adhesive tracks with solution gradients and therefore allows one to determine the relative importance of guidance cues.

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as ...

creating-adhesive-soluble-gradients-for-imaging-cell-migration-with

Research

JoVE Journal

Bioengineering

12.5K Views.  The University of New South Wales. A method for the assembly of adhesive and soluble gradients in a microscopy chamber for live cell migration studies is described. The engineered environment combines antifouling surfaces and adhesive tracks with solution gradients and therefore allows one to determine the relative importance of guidance cues.

Video: Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy

Insertion of Flexible Neural Probes Using Rigid Stiffeners Attached with Biodissolvable Adhesive

Microelectrode arrays for neural interface devices that are made of biocompatible thin-film polymer are expected to have extended functional lifetime because the flexible material may minimize adverse tissue response caused by micromotion. However, their flexibility prevents them from being accurately inserted into neural tissue. This article demonstrates a method to temporarily attach a flexible microelectrode probe to a rigid stiffener using biodissolvable polyethylene glycol (PEG) to facilitate precise, surgical insertion of the probe. A unique stiffener design allows for uniform distribution of the PEG adhesive along the length of the probe. Flip-chip bonding, a common tool used in microelectronics packaging, enables accurate and repeatable alignment and attachment of the probe to the stiffener. The probe and stiffener are surgically implanted together, then the PEG is allowed to dissolve so that the stiffener can be extracted leaving the probe in place. Finally, an in vitro test method is used to evaluate stiffener extraction in an agarose gel model of brain tissue. This approach to implantation has proven particularly advantageous for longer flexible probes (&gt;3 mm). It also provides a feasible method to implant dual-sided flexible probes. To date, the technique has been used to obtain various in vivo recording data from the rat cortex.

Insertion of flexible neural microelectrode probes is enabled by attaching probes to rigid stiffeners with polyethylene glycol (PEG). A unique assembly process ensures uniform and repeatable attachment. After insertion into tissue, the PEG dissolves and the stiffener is extracted. An in vitro test method evaluates the technique in agarose gel.

Microelectrode  arrays for neural interface devices that are made of biocompatible thin-film  polymer are expected to have extended functional lifetime ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM)

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and sensitivity in cells because the resolution of conventional light microscopy is diffraction-limited to ~250 nm. By combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM), PPIs can be visualized in cells with single molecule sensitivity and nanometer spatial resolution. BiFC is a commonly used technique for visualizing PPIs with fluorescence contrast, which involves splitting of a fluorescent protein into two non-fluorescent fragments. PALM is a recent superresolution microscopy technique for imaging biological samples at the nanometer and single molecule scales, which uses phototransformable fluorescent probes such as photoactivatable fluorescent proteins (PA-FPs). BiFC-PALM was demonstrated by splitting PAmCherry1, a PA-FP compatible with PALM, for its monomeric nature, good single molecule brightness, high contrast ratio, and utility for stoichiometry measurements. When split between amino acids 159 and 160, PAmCherry1 can be made into a BiFC probe that reconstitutes efficiently at 37 &#176;C with high specificity to PPIs and low non-specific reconstitution. Ras-Raf interaction is used as an example to show how BiFC-PALM helps to probe interactions at the nanometer scale and with single molecule resolution. Their diffusion can also be tracked in live cells using single molecule tracking (smt-) PALM. In this protocol, factors to consider when designing the fusion proteins for BiFC-PALM are discussed, sample preparation, image acquisition, and data analysis steps are explained, and a few exemplary results are showcased. Providing high spatial resolution, specificity, and sensitivity, BiFC-PALM is a useful tool for studying PPIs in intact biological samples.

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation BiFC-PALM

Protein-protein interactions are visualized in cells with nanometer spatial resolution by combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM). Described here is the use of BiFC-PALM for imaging Ras-Raf interactions in U2OS cells for visualizing the nanoscale clustering and diffusion of individual Ras-Raf complexes.

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and ...

Polydimethylsiloxane-polycarbonate Microfluidic Devices for Cell Migration Studies Under Perpendicular Chemical and Oxygen Gradients

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under combinations of chemical and oxygen gradients. Both chemical and oxygen gradients can greatly affect cell migration in vivo; however, due to technical limitations, very little research has been performed to investigate their effects in vitro. The device developed in this research takes advantage of a series of serpentine-shaped channels to generate the desired chemical gradients and exploits a spatially confined chemical reaction method for oxygen gradient generation. The directions of the chemical and oxygen gradients are perpendicular to each other to enable straightforward migration result interpretation. In order to efficiently generate the oxygen gradients with minimal chemical consumption, the embedded PC thin film is utilized as a gas diffusion barrier. The developed microfluidic device can be actuated by syringe pumps and placed into a conventional cell incubator during cell migration experiments to allow for setup simplification and optimized cell culture conditions. In cell experiments, we used the device to study migrations of adenocarcinomic human alveolar basal epithelial cells, A549, under combinations of chemokine (stromal cell-derived factor, SDF-1&#945;) and oxygen gradients. The experimental results show that the device can stably generate perpendicular chemokine and oxygen gradients and is compatible with cells. The migration study results indicate that oxygen gradients may play an essential role in guiding cell migration, and cellular behavior under combinations of gradients cannot be predicted from those under single gradients. The device provides a powerful and practical tool for researchers to study interactions between chemical and oxygen gradients in cell culture, which can promote better cell migration studies in more in vivo-like microenvironments.

The control of chemical and oxygen gradients is essential for cell cultures. This paper reports a polydimethylsiloxane-polycarbonate (PDMS-PC) microfluidic device capable of reliably generating combinations of chemical and oxygen gradients for cell migration studies, which can be practically utilized in biological labs without sophisticated instrumentation.

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under ...

Conducting Multiple Imaging Modes with One Fluorescence Microscope

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to conventional epi-fluorescence microscopy, various imaging techniques have been developed to achieve specific experimental goals. Some of the widely used techniques include single-molecule fluorescence resonance energy transfer (smFRET), which can report conformational changes and molecular interactions with angstrom resolution, and single-molecule detection-based super-resolution (SR) imaging, which can enhance the spatial resolution approximately ten&#160;to twentyfold compared to diffraction-limited microscopy. Here we present a customer-designed integrated system, which merges multiple imaging methods in one microscope, including conventional epi-fluorescent imaging, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging. Different imaging methods can be achieved easily and reproducibly by switching optical elements. This set-up is easy to adopt by any research laboratory in biological sciences with a need for routine and diverse imaging experiments at a reduced cost and space relative to building separate microscopes for individual purposes.

Here we present a practical guide of building an integrated microscopy system, which merges conventional epi-fluorescent imaging, single-molecule detection-based super-resolution imaging, and multi-color single-molecule detection, including single-molecule fluorescence resonance energy transfer imaging, into one set-up in a cost-efficient way.

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to ...

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to visualize these highly dynamic events, two complementary light microscopy techniques that allow fast imaging of live samples were combined: spinning disk microscopy (SD) for fast and high-resolution volume recording and total internal reflection fluorescence (TIRF) microscopy for precise localization and visualization of the plasma membrane. A comprehensive and complete imaging protocol will be shown for guiding through sample preparation, microscope calibration, image formation and acquisition, resulting in multi-color SD-TIRF live imaging series with high spatio-temporal resolution. All necessary image post-processing steps to generate multi-dimensional live imaging datasets, i.e. registration and combination of the individual channels, are provided in a self-written macro for the open source software ImageJ. The imaging of fluorescent proteins during initiation and maturation of adhesion complexes, as well as the formation of the actin cytoskeletal network, was used as a proof of principle for this novel approach. The combination of high resolution 3D microscopy and TIRF provided a detailed description of these complex processes within the cellular environment and, at the same time, precise localization of the membrane-associated molecules detected with a high signal-to-background ratio.

An advanced microscope that permit fast and high-resolution imaging of both, the isolated plasma membrane and the surrounding intracellular volume, will be presented. The integration of spinning disk and total internal reflection fluorescence microscopy in one setup allows live imaging experiments at high acquisition rates up to 3.5 s per image stack.

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to ...

3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and chemical complexity of living tissues and mimic in vivo tissue functions. Recent advances in microfabrication technologies have facilitated the development of 3D in vitro environments in which cells can be integrated into a well-defined extracellular matrix (ECM) and a defined set of soluble or matrix associated biomolecules. However, technological barriers have limited their widespread use in research laboratories. Here, we describe a method to construct simple devices for 3D culture and experimentation with cells and multicellular organoids in 3D microenvironments with a defined chemoattractant gradient. We illustrate the use of this platform for analysis of the response of epithelial cells and organoids to gradients of growth factors, such as epidermal growth factor (EGF). EGF gradients were stable in the devices for several days leading to directed branch formation in breast organoids. This analysis allowed us to conclude that collective gradient sensing by groups of cells is more sensitive vs. single cells. We also describe the fabrication method, which does not require photolithography facilities nor advanced soft lithography techniques. This method will be helpful to study 3D cellular behaviors in the context of the analysis of development and pathological states, including cancer.

We describe a method to construct devices for 3D culture and experimentation with cells and multicellular organoids. This device allows analysis of cellular responses to soluble signals in 3D microenvironments with defined chemoattractant gradients. Organoids are better than single cells at detection of weak noisy inputs.

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and ...

Traction Microscopy Integrated with Microfluidics for Chemotactic Collective Migration

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical gradient, known as chemotaxis, plays an important role in development, the immune response, wound healing, and cancer metastasis. While chemotaxis modulates the migration of single cells as well as collections of cells in vivo, in vitro research focuses on single-cell chemotaxis, partly due to the lack of the proper experimental tools. To fill that gap, described here is a unique experimental system that combines microfluidics and micropatterning to demonstrate the effects of chemical gradients on collective cell migration. Furthermore, traction microscopy and monolayer stress microscopy are incorporated into the system to characterize changes in cellular force on the substrate as well as between neighboring cells. As proof-of-concept, the migration of micropatterned circular islands of Madin-Darby canine kidney (MDCK) cells is tested under a gradient of hepatocyte growth factor (HGF), a known scattering factor. It is found that cells located near the higher concentration of HGF migrate faster than those on the opposite side within a cell island. Within the same island, cellular traction is similar on both sides, but intercellular stress is much lower on the side of higher HGF concentration. This novel experimental system can provide new opportunities to studying the mechanics of chemotactic migration by cellular collectives.

Collective cell migration in development, wound healing, and cancer metastasis is often guided by the gradients of growth factors or signaling molecules. Described here is an experimental system combining traction microscopy with a microfluidic system and a demonstration of how to quantify the mechanics of collective migration under biochemical gradient.

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical ...