Funding support provided by NIH grant HL89764, NSF grants 0546012, 0730503 and 0725528.

A. Biotinylation of DNA
Plasmid DNA were covalently biotinylated with guanine-specific biotin labels as described by the manufacturer (Mirus Bio, Madison, WI) but scaled to have ~1-2 biotin labels per DNA. Plasmid DNA (pEGFP-C1, 4.9 kb, Clontech, Mountain View, CA) was labeled in this protocol.
<ol><li>Dissolve desired amount of pDNA into TE buffer of DNase-free and RNase-free (molecular biology-grade quality) water to make a 1&mu;g/&mu;L DNA solution.</li>
 <li>Conduct the labeling reaction using the following reaction mixtures. Add the LabelIT reagent last.</li>
</ol>
 For 100&mu;g DNA reaction:
<table border="1">
<tbody>
 <tr>
 <td>
 DNase-free and RNase-free water </td>
 <td>75 &mu;L</td>
 </tr>
 <tr>
 <td>10X Labeling Buffer A</td>
 <td>20 &mu;L</td>
 </tr>
 <tr>
 <td>1&mu;g/&mu;L DNA</td>
 <td>100 &mu;L</td>
 </tr>
 <tr>
 <td>LabelIT reagent</td>
 <td>5 &mu;L</td>
 </tr>
 <tr>
 <td>Total Volume</td>
 <td>200 &mu;L</td>
 </tr>
</tbody>
</table>
<ol start="3">
 <li>Incubate the reaction at 37 &deg;C for 1 hour.</li>
 <li>Purify the labeled sample by ethanol or isopropanol precipitation following standard protocols.</li>
</ol>
Note: Gel filtration based columns may lead to high UV absorbance or fluorescence background, which may affect the DNA quantification or fluorescence characterization. 
 Note: The level of biotinylation may be determined by HABA-based tests.
 B. Labeling of the Cy5-Cationic Polymer
Chitosan (390 kDa, 83.5% deacetylated, Vanson, Redmond, WA) was used as a model cationic polymer in this study. The free primary amines on the chitosan polymer backbone were labeled with Cy5-NHS (Amersham Biosciences, Piscataway, NJ).
<ol>
 <li>To facilitate complete conjugation of Cy5 dye, calculate the required amount of Cy5-NHS such that the molar ratio of Cy5 : primary amines is 1 : 200.</li>
 <li>Adjust pH of the chitosan solution (in 25mM acetate buffer) to ~6.5 by addition of NaOH. Note that the NHS reaction is more efficient at basic pH, but the solubility of chitosan here limits the working pH range.</li>
 <li>While stirring, slowly add the calculated amount of Cy5-NHS (1 mg/ml DMSO) to the chitosan solution in a drop-by-drop manner.</li>
 <li>Agitate the mixture in the dark at room temperature overnight.</li>
 <li>To purify, dialyze with 10k MWCO Slide-a-Lyzer (Pierce) for 2 hr against 1% acetate buffer at room temp in the dark.</li>
 <li>Replace buffer and dialyze another 2 hr at room temp in the dark.</li>
 <li>Replace buffer and dialyze overnight at 4 &deg;C in the dark.</li>
 <li>Store purified labeled polymer at -20 &deg;C.</li></ol>
Note: In this study, a standard curve is constructed by measuring the emission intensity of Cy5-NHS ester at 670 nm. Characterize the labeling density by measuring the obtained emission at 670 nm from Cy5-labeled chitosan in the standard curve. Absorbance may also be used to determine the labeling efficiency but was not performed here.
C. Preparation of QD-labeled DNA and Cy5-Polymer
The molar ratio of pDNA to QD was kept in excess (pDNA : QD &asymp; 1 : 2) to ensure complete conjugation of QDs to pDNA. The number of QDs labeled onto each pDNA can be estimated through TEM imaging or other equivalent facilities. In our study, the number of QDs per pDNA is estimated to be ~1-3 by TEM and single molecule spectroscopy.1 Use Millipore Milli-Q gradient water (&gt;18.0 MW, 0.2um filtered) during the preparation.
<ol> 
 <li>Calculate the required amount of chitosan for 10&mu;g pDNA according to desired N/P ratio, the theoretical ratio of protonated amines in the chitosan solution to the negative phosphates in the DNA solution.</li>
 <li>Add streptavidin-functionalized 605QDs (Qdot 605 ITK, Invitrogen, Carlsbad, CA) into the biotinylated pDNA solution.</li>
 <li>Incubate the solution at room temperature in the dark for 15 min.</li>
 <li>Add the QD-labeled DNA into 50 mM sodium sulfate solution to make the final volume 200&mu;L.</li>
 <li>Dilute Cy5-chitosan, according to desired N/P ratio, with Milli-Q water to make the final volume 200&mu;L.</li>
 </ol>
 Note: Keep the reaction in dark to prevent possible photobleaching. 
 Important: Be cautious to use the Qdot 605 ITK&trade; streptavidin conjugate (the ITK series), as Quantum dots in this catalog are designed for the purpose of FRET. The regular Qdot series are conjugated with a PEG layer to prevent non-specific binding, especially for cellular labeling. However, this additional coating enlarges the donor-acceptor distance, resulting in reduced energy transfer efficiency.
 D. Fabrication of the SU-8 Masters Using Standard Photolithography
<ol>
 <li>Si wafer is piranha cleaned and baked at 200 &deg;C for 5 min.</li>
 <li>For the designated master thickness of 25&mu;m, spin coat the negative photoresist (SU-8 2025, Microchem, Newton, MA) on Si wafer at 2000 rpm for 30 sec.</li>
 <li>Soft bake the wafer on a hotplate with a ramp of 65 &deg;C/hr to 95 &deg;C.</li>
 <li>Expose to UV light (365nm) for 250mJ/cm2 through a mask film (CAD/Art Services, Bandon, OR) containing the design of microchannels.</li>
 <li>Post-exposure bake the wafer on a hotplate with a ramp of 65 &deg;C/hr to 95 &deg;C.</li>
 <li>Develop the wafer using SU-8 photoresist developer.</li>
 <li>The patterned wafer is hard baked on a hotplate with a ramp of 65 &deg;C/hr to 200 &deg;C. Maintain the wafer at 200 &deg;C for at least 5 hours, then gradually cool the wafer down to room temperature.</li>
 </ol>
 Important: Gradual ramping during the SU-8 master baking process is necessary, otherwise the SU-8 structure may detached from the silicon wafer or cracks on the SU-8 structure may be induced by stress-release.
 E. Replica Molding of PDMS from the masters and Bonding to the Cover Glass
<ol>
 <li>The SU-8 master is placed in a weighing boat.</li>
 <li>Mix silicone elastomer and curing agent (Poly(dimethylsiloxane), PDMS, Sylgard 184, Dow Corning, Midland, MI)&nbsp; in a 10 : 1 ratio.</li>
 <li>Pour the PDMS mixture onto the SU-8 master and leave the weighing boat in a vacuum desiccator to remove bubbles.</li>
 <li>Cure the PDMS at 65 &deg;C for 1-2 hours.</li>
 <li>Peel the PDMS strip from the Si master mold.</li>
 <li>Punch channel inlets and outlets of the fluidic device.</li>
 <li>Clean the PDMS strip and cover glass with ethanol and then air-dry.</li>
 <li>Treat the cleaned PDMS strip and cover glass with oxygen plasma (20W for 1min).</li>
 <li>Immediately bond the PDMS strip with cover glass.</li>
 <li>Leave the bonded microfluidic chip in the oven at 95 &deg;C for overnight.</li>
</ol>
Important: Plasma treatment and overnight baking are essential to enhanced bonding strength.
F. Monitor the formation of DNA Nanocomplexes In the Microfluidic Device
<ol>
 <li>Fill the microfluidic channel with water (to ensure there is no bubbles within the microfluidic channel), before loading the reagents to ensure smooth flow during the experiment.</li>
 <li>Load the QD-labeled DNA and Cy5-labeled chitosan solutions into two individual glass syringes, through the tubing described in the video.</li>
 <li>Connect the tubing with the two inlets of microfluidic devices. Be cautious not to introduce any air during the process. Set the flow rate at 20nL/min (PHD-2000 syringe pump, Holliston, MA), under laminar flow conditions.</li>
 <li>Check the microchannels under the microscope.</li>
 <li>When the flow is stable (~15 to 20 minutes), QD-mediated FRET should be observed in the center of the channel.</li>
 <li>Take fluorescence pictures (Cooled CCD, Qimaging, BC, Canada) at different locations along the channel.</li>
 <li>Analyze the fluorescence images with ImageJ and OriginLab.</li>
</ol>
<img src="/files/ftp_upload/1432/1432fig1.jpg" alt="Figure 1" /> Figure 1. QD-FRET provides a sensitive indication of the onset of DNA Nanocomplexes self-assembly
<ol style="list-style-type:lower-alpha;">
 <li>Quantum dot-mediated fluorescence resonance energy transfer (QD-FRET) can provide a quantitative and highly sensitive indication of polyplex stability in either extra- or intra-cellular environments, allowing for unambiguous detection of the onset of interactions between DNA and the gene carrier. The FRET pair, 605QD and Cy5, was chosen based on maximizing spectral overlap between the donor and acceptor and minimizing potential cross-talk.&nbsp; For this pair, the F&ouml;rster distance is 69.4&Aring;.3</li>
 <li>Self-assembly of the QD-FRET DNA nanocomplexes. Anionic plasmid DNA (pDNA) and the cationic gene carrier were labeled with QD (energy donor) and Cy5 (energy acceptor), respectively. QD-FRET nanocomplexes were formed through electrostatic complex coacervation. Upon excitation at 488 nm, QD-FRET-mediated Cy5 emission indicated formation of a compact and intact nanocomplex. The residence time (tR) can be calculated according to the distance (x) which measures from where the two streams meet to the position of reaction under investigation, and the mean flow speed (v). Due to the nature of laminar flow, mixing only takes place at the interface (center of each image), allowing precise calculation of mass transport as a function of tR. Temporal resolution can be adjusted by varying the applied flow rates. (Inset) FRET-mediated signal was observed immediately at the interface when the two streams met, indicating that binding was rapid, occurring within a few milliseconds according to the applied flow rates. Scale Bar: 100&mu;m.</li>
</ol>

<ol>
	<li>Ho, Y. P., Chen, H. H., Leong, K. W., Wang, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluating+the+intracellular+stability+and+unpacking+of+DNA+nanocomplexes+by+quantum+dots-FRET.">Evaluating the intracellular stability and unpacking of DNA nanocomplexes by quantum dots-FRET.</a> J Control Release. 116, 83-89 (2006).</li><li>Chen, H. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+comparison+of+intracellular+unpacking+kinetics+of+polyplexes+by+a+model+constructed+from+quantum+dot-FRET.">Quantitative comparison of intracellular unpacking kinetics of polyplexes by a model constructed from quantum dot-FRET.</a> Mol. Ther. 16, 324-332 (2008).</li><li>Zhang, C. Y., Yeh, H. C., Kuroki, M., Wang, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-Quantum-Dot-Based+DNA+Nanosensor.">Single-Quantum-Dot-Based DNA Nanosensor.</a> Nat Mat. 4, 826-831 (2005).</li></ol>

The authors have nothing to disclose.

<ul><li>Significance of our work:
 <ol>
 <li>This is the first attempt to monitor polymeric DNA nanocomplex self-assembly kinetics in real-time (millisecond resolution) through QD-FRET responses within a simple microfluidic chip. </li>
 <li>QD-mediated FRET provides a highly sensitive and quantitative indication of the onset of molecular interactions and throughout the self-assembly process, whereas microfluidics offers a well-controlled microenvironment to spatially analyze the process during the DNA nanocomplex synthesis. </li>
 <li>The integration of microfluidics and nanophotonics suggests a new and interesting approach to investigate any type of complexation reactions.</li>
 <li>The resulting QD-FRET polymeric DNA nanocomplexes could be readily applied for establishing structure-function relationships.1,2</li>
 </ol>
</li></ul>

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>SU-8</td>
<td>&nbsp;</td>
<td>MicroChem</td>
<td>SU-8 2025</td>
<td>&nbsp;</td>
<tr>
<td>PDMS</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<tr>
<td>Plasmid DNA</td>
<td>&nbsp;</td>
<td>Clontech</td>
<td>pEGFP-C1</td>
<td>4.9 kb, MW = 3.3 x 106</td>
</tr>
<tr>
<td>LabelIT Biotin Labeling Kit</td>
<td>&nbsp;</td>
<td>Mirus Bio LLC</td>
<td>MIR 3400</td>
<td>Standard protocol yields labeling efficiency of approximately one label every 20-60 bp of double-stranded DNA, The density of labeling was adjusted in this work.</td>
</tr>
<tr>
<td>Streptavidin 605QD</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>Qdot&reg; 605 ITK&trade; Streptavidin Conjugate</td>
<td>&nbsp;</td>
<tr>
<td>Cy5-NHS Ester</td>
<td>&nbsp;</td>
<td>Amersham Biosciences</td>
<td>PA15101</td>
<td>&nbsp;</td>
<tr>
<td>Chitosan</td>
<td>&nbsp;</td>
<td>Vanson</td>
<td>390 kDa </td>
<td>83.5% deacetylated</td>
</tr>
<tr>
<td>Cover Glass</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>12-545C</td>
<td>No. 1; Size: 40 x 22mm</td>
</tr>
<tr>
<td>Gastight Glass Syringe</td>
<td>&nbsp;</td>
<td>Hamilton</td>
<td>TLL series</td>
<td>50&mu;L to 500&mu;L depending on sample volume</td>
</tr>
<tr>
<td>Tygon Tubing</td>
<td>&nbsp;</td>
<td>SmallParts</td>
<td>0.02 ID, 100ft</td>
<td>Tygon Tubes Microbore, 0.02 ID, 100ft</td>
</tr>
<tr>
<td>Connector</td>
<td>&nbsp;</td>
<td>SmallParts</td>
<td>HTX-23R</td>
<td>Customized in length of 0.750"</td>
</tr>
<tr>
<td>Syringe Pump</td>
<td>&nbsp;</td>
<td>Harvard Apparatus</td>
<td>PHD-2000</td>
<td>&nbsp;</td>
<tr>
<td>CCD</td>
<td>&nbsp;</td>
<td>Qimaging</td>
<td>Intensified Retiga Cooled</td>
<td>&nbsp;</td>
<tr>
<td>Microscope</td>
<td>&nbsp;</td>
<td>Olympus</td>
<td>BX-51</td>
<td>100W mercury arc lamp</td>
</tr>
<tr>
<td>ImageJ</td>
<td>&nbsp;</td>
<td>NIH</td>
<td>v1.36b</td>
<td>http://rsb.info.nih.gov/ij</td>
</tr>
<tr>
<td>Origin Pro8</td>
<td>&nbsp;</td>
<td>OriginLab</td>
<td>Student Version</td>
<td>&nbsp;</td>
<tr>
<td>Microscope Filter sets</td>
<td>&nbsp;</td>
<td>Omega Optical</td>
<td>475AF40</td>
<td>Excitation filter in both channels</td>
</tr>
<tr>
<td>Microscope Filter sets</td>
<td>&nbsp;</td>
<td>Omega Optical</td>
<td>595AF60</td>
<td>Emission filter in 605QD channel</td>
</tr>
<tr>
<td>Microscope Filter sets</td>
<td>&nbsp;</td>
<td>Omega Optical</td>
<td>670DF40</td>
<td>Emission filter in QD-FRET channel</td>
</tr>
<tr>
<td>Microscope Filter sets</td>
<td>&nbsp;</td>
<td>Omega Optical</td>
<td>500 DRLP</td>
<td>Long pass dichroic in 605QD channel</td>
</tr>
<tr>
<td>Microscope Filter sets</td>
<td>&nbsp;</td>
<td>Omega Optical</td>
<td>595DRLP</td>
<td>Long pass dichroic in QD-FRET channel</td>
</tr>		</tbody>
		</table>
		</div>

combining qd-fret and microfluidics to monitor dna nanocomplex self-assembly in real-time

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically functional inside the cell, nucleic acid-based therapeutics require an efficient intracellular delivery system. One widely adopted approach is to complex DNA with a gene carrier to form nanocomplexes via electrostatic self-assembly, facilitating cellular uptake of DNA while protecting it against degradation. The challenge lies in the rational design of efficient gene carriers, since premature dissociation or overly stable binding would be detrimental to the cellular uptake and therapeutic efficacy. Nanocomplexes synthesized by bulk mixing showed a diverse range of intracellular unpacking and trafficking behavior, which was attributed to the heterogeneity in size and stability of nanocomplexes. Such heterogeneity hinders the accurate assessment of the self-assembly kinetics and adds to the difficulty in correlating their physical properties to transfection efficiencies or bioactivities. We present a novel convergence of nanophotonics (i.e. QD-FRET) and microfluidics to characterize the real-time kinetics of the nanocomplex self-assembly under laminar flow. QD-FRET provides a highly sensitive indication of the onset of molecular interactions and quantitative measure throughout the synthesis process, whereas microfluidics offers a well-controlled microenvironment to spatially analyze the process with high temporal resolution (~milliseconds). For the model system of polymeric nanocomplexes, two distinct stages in the self-assembly process were captured by this analytic platform. The kinetic aspect of the self-assembly process obtained at the microscale would be particularly valuable for microreactor-based reactions which are relevant to many micro- and nano-scale applications. Further, nanocomplexes may be customized through proper design of microfludic devices, and the resulting QD-FRET polymeric DNA nanocomplexes could be readily applied for establishing structure-function relationships. 

Watch this Scientific Journal Video about Combining QD-FRET and Microfluidics to Monitor DNA Nanocomplex Self-Assembly in Real-Time at JoVE.com

Combining QD-FRET and Microfluidics to Monitor DNA Nanocomplex Self-Assembly in Real-Time

We present a novel and powerful integration of nanophotonics (QD-FRET) and microfluidics to investigate the formation of polyelectrolyte polyplexes, which is expected to provide better control and synthesis of uniform and customizable polyplexes for future nucleic acid-based therapeutics.

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically ...

combining-qd-fret-microfluidics-to-monitor-dna-nanocomplex-self

Research

JoVE Journal

Biology

11.0K Views.  Johns Hopkins University. We present a novel and powerful integration of nanophotonics (QD-FRET) and microfluidics to investigate the formation of polyelectrolyte polyplexes, which is expected to provide better control and synthesis of uniform and customizable polyplexes for future nucleic acid-based therapeutics.

Video: Combining QD-FRET and Microfluidics to Monitor DNA Nanocomplex Self-Assembly in Real-Time

Chemotactic Response of Marine Micro-Organisms to Micro-Scale Nutrient Layers

The degree to which planktonic microbes can exploit microscale resource patches will have considerable implications for oceanic trophodynamics and biogeochemical flux. However, to take advantage of nutrient patches in the ocean, swimming microbes must overcome the influences of physical forces including molecular diffusion and turbulent shear, which will limit the availability of patches and the ability of bacteria to locate them. Until recently, methodological limitations have precluded direct examinations of microbial behaviour within patchy habitats and realistic small-scale flow conditions. Hence, much of our current knowledge regarding microbial behaviour in the ocean has been procured from theoretical predictions. To obtain new information on microbial foraging behaviour in the ocean we have applied soft lithographic fabrication techniques to develop 2 microfluidic devices, which we have used to create (i) microscale nutrient patches with dimensions and diffusive characteristics relevant to oceanic processes and (ii) microscale vortices, with shear rates corresponding to those expected in the ocean. These microfluidic devices have permitted a first direct examination of microbial swimming and chemotactic behaviour within a heterogeneous and dynamic seascape. The combined use of epifluorescence and phase contrast microscopy allow direct examinations of the physical dimensions and diffusive characteristics of nutrient patches, while observing the population-level aggregative response, in addition to the swimming behaviour of individual microbes. These experiments have revealed that some species of phytoplankton, heterotrophic bacteria and phagotrophic protists are adept at locating and exploiting diffusing microscale resource patches within very short time frames. We have also shown that up to moderate shear rates, marine bacteria are able to  fight the flow  and swim through their environment at their own accord. However, beyond a threshold high shear level, bacteria are aligned in the shear flow and are less capable of swimming without disturbance from the flow. Microfluidics represents a novel and inexpensive approach for studying aquatic microbial ecology, and due to its suitability for accurately creating realistic flow fields and substrate gradients at the microscale, is ideally applicable to examinations of microbial behaviour at the smallest scales of interaction.  We therefore suggest that microfluidics represents a valuable tool for obtaining a better understanding of the ecology of microorganisms in the ocean.

The fabrication of microfluidic channels and their implementation in experiments for studying the chemotactic foraging behaviour of marine microbes within a patchy nutrient seascape and the swimming behaviour of bacteria within shear flow are described.

The degree to which planktonic microbes can exploit microscale resource patches will have considerable implications for oceanic trophodynamics and ...

Studies of Bacterial Chemotaxis Using Microfluidics - Interview

Applying Microfluidics to Electrophysiology

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the microfluidic brain slice device is discussed including how the device docks to standard perfusion chambers and the technique of passive pumping which is used to deliver boluses of neuromodulators to the brain slice.  By simplifying the device design, we are able to achieve a practical solution to the current unmet electrophysiology need of applying multiple neuromodulators across multiple regions of the brain slice.  This is achieved by substituting the standard coverglass substrate of the perfusion chamber with a thin microfluidic device bonded to the coverglass substrate.  This was then attached to the perfusion chamber and small holes connect the open-well of the perfusion chamber to the microfluidic channels buried within the microfluidic substrate.  These microfluidic channels are interfaced with ports drilled into the edge of the perfusion chamber to access and deliver stimulants.  This project represents how the field of microfluidics is transitioning away from proof-of concept device demonstrations and into practical solutions for unmet experimental and clinical needs.

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the ...

Digital Microfluidics for Automated Proteomic Processing

Clinical proteomics has emerged as an important new discipline, promising the discovery of biomarkers that will be useful for early diagnosis and prognosis of disease. While clinical proteomic methods vary widely, a common characteristic is the need for (i) extraction of proteins from extremely heterogeneous fluids (i.e. serum, whole blood, etc.) and (ii) extensive biochemical processing prior to analysis. Here, we report a new digital microfluidics (DMF) based method integrating several processing steps used in clinical proteomics. This includes protein extraction, resolubilization, reduction, alkylation and enzymatic digestion. Digital microfluidics is a microscale fluid-handling technique in which nanoliter-microliter sized droplets are manipulated on an open surface. Droplets are positioned on top of an array of electrodes that are coated by a dielectric layer - when an electrical potential is applied to the droplet, charges accumulate on either side of the dielectric. The charges serve as electrostatic handles that can be used to control droplet position, and by biasing a sequence of electrodes in series, droplets can be made to dispense, move, merge, mix, and split on the surface. Therefore, DMF is a natural fit for carrying rapid, sequential, multistep, miniaturized automated biochemical assays. This represents a significant advance over conventional methods (relying on manual pipetting or robots), and has the potential to be a useful new tool in clinical proteomics.
Mais J. Jebrail, Vivienne N. Luk, and Steve C. C. Shih contributed equally to this work.
Sergio L. S. Freire's current address is at the University of the Sciences in Philadelphia located at 600 South 43rd Street, Philadelphia, PA 19104.

Digital Microfluidics is a technique characterized by the manipulation of discrete droplets (~nL - mL) on an array of electrodes by the application of electrical fields. It is well-suited for carrying out rapid, sequential, miniaturized automated biochemical assays. Here, we report a platform capable of automating several proteomic processing steps.

Clinical proteomics has emerged as an important new discipline, promising the discovery of biomarkers that will be useful for early diagnosis and ...

Determining Genetic Expression Profiles in C. elegans Using Microarray and Real-time PCR

Synapses are composed of a presynaptic active zone in the signaling cell and a postsynaptic terminal in the target cell. In the case of chemical synapses, messages are carried by neurotransmitters released from presynaptic terminals and received by receptors on postsynaptic cells. Our previous research in Caenorhabditis elegans has shown that VSM-1 negatively regulates exocytosis. Additionally, analysis of synapses in vsm-1 mutants showed that animals lacking a fully functional VSM-1 have increased synaptic connectivity. Based on these preliminary findings, we hypothesized that C. elegans VSM-1 may play a crucial role in synaptogenesis. To test this hypothesis, double-labeled microarray analysis was performed, and gene expression profiles were determined. First, total RNA was isolated, reversely transcribed to cDNA, and hybridized to the DNA microarrays. Then, in-silico analysis of fluorescent probe hybridization revealed significant induction of many genes coding for members of the major sperm protein family (MSP) in mutants with enhanced synaptogenesis. MSPs are the major component of sperm in C. elegans and appear to signal nematode oocyte maturation and ovulation . In fruit flies, Chai and colleagues 1 demonstrated that MSP-like molecules regulate presynaptic bouton number and size at the neuromuscular junction. Moreover, analysis performed by Tsuda and coworkers 2 suggested that MSPs may act as ligands for Eph receptors and trigger receptor tyrosine kinase signaling cascades. Lastly, real time PCR analysis corroborated that the gene coding for MSP-32 is induced in vsm-1(ok1468) mutants. Taken together, research performed by our laboratory has shown that vsm-1 mutants have a significant increase in synaptic density, which could be mediated by MSP-32 signaling.

Determining Genetic Expression Profiles in C. elegans Using Microarray and Real-time PCR

Microarray analysis was conducted to determine genetic expression profiles in C. elegans, and real-time PCR was used to validate and quantify microarray data.

Synapses are composed of a presynaptic active zone in the signaling cell and a postsynaptic terminal in the target cell. In the case of chemical synapses, ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix containing polysaccharides that include cellulose microfibrils that form both crystalline structures and cellulose chains of amorphous organization. The orientation of the cellulose fibers and their concentrations dictate the mechanical properties of the cell. Several methods are used to determine the levels of crystalline cellulose, each bringing both advantages and limitations. Some can distinguish the proportion of crystalline regions within the total cellulose. However, they are limited to whole-organ analyses that are deficient in spatiotemporal information. Others relying on live imaging, are limited by the use of imprecise dyes. Here, we report a sensitive polarized light-based system for specific quantification of relative light retardance, representing crystalline cellulose accumulation in cross sections of Arabidopsis thaliana roots. In this method, the cellular resolution and anatomical data are maintained, enabling direct comparisons between the different tissues composing the growing root. This approach opens a new analytical dimension, shedding light on the link between cell wall composition, cellular behavior and whole-organ growth.

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix ...

Detecting and Characterizing Protein Self-Assembly In Vivo by Flow Cytometry

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from low-sensitivity, indirect read-outs, limited throughput, and/or population-level rather than single-cell resolution. We designed a flow cytometry-based single methodology that addresses all of these limitations: Distributed Amphifluoric FRET or DAmFRET. DAmFRET detects and quantifies protein self-assemblies by sensitized emission FRET in vivo, enables deployment across model systems-from yeast to human cells-and achieves sensitive, single-cell, high-throughput read-outs irrespective of protein localization or solubility.

This article describes a FRET-based flow cytometry protocol to quantify protein self-assembly in both S. cerevisiae and HEK293T cells.

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from ...

Wide-Field, Real-Time Imaging of Local and Systemic Wound Signals in Arabidopsis

Plants respond to mechanical stresses such as wounding and herbivory by inducing defense responses both in the damaged and in the distal undamaged parts. Upon wounding of a leaf, an increase in cytosolic calcium ion concentration (Ca2+ signal) occurs at the wound site. This signal is rapidly transmitted to undamaged leaves, where defense responses are activated. Our recent research revealed that glutamate leaking from the wounded cells of the leaf into the apoplast around them serves as a wound signal. This glutamate activates glutamate receptor-like Ca2+ permeable channels, which then leads to long-distance Ca2+ signal propagation throughout the plant. The spatial and temporal characteristics of these events can be captured with real-time imaging of living plants expressing genetically encoded fluorescent biosensors. Here we introduce a plant-wide, real-time imaging method to monitor the dynamics of both the Ca2+ signals and changes in apoplastic glutamate that occur in response to wounding. This approach uses a wide-field fluorescence microscope and transgenic Arabidopsis plants expressing Green Fluorescent Protein (GFP)-based Ca2+ and glutamate biosensors. In addition, we present methodology to easily elicit wound-induced, glutamate-triggered rapid and long-distance Ca2+ signal propagation. This protocol can also be applied to studies on other plant stresses to help investigate how plant systemic signaling might be involved in their signaling and response networks.

Wide-Field, Real-Time Imaging of Local and Systemic Wound Signals in Arabidopsis

Extracellular glutamate-triggered systemic calcium signaling is critical for the induction of plant defense responses to mechanical wounding and herbivore attack in plants. This article describes a method to visualize the spatial and temporal dynamics of both these factors using Arabidopsis thaliana plants expressing calcium- and glutamate-sensitive fluorescent biosensors.

Plants respond to mechanical stresses such as wounding and herbivory by inducing defense responses both in the damaged and in the distal undamaged parts. ...