We thank Philipp Denninger for help with video preparation and Bhavna Chaudhuri for providing plant lines expressing FRET sensors. This work was supported by grants from the National Science Foundation (MCB 1021677), the Department of Energy (DE-FG02-04ER15542) to W.B.F, the National Institutes of Health, and the Howard Hughes Medical Institute to S.R.Q. G.G. was supported by an EMBO long-term fellowship. M.M. was supported by the Alexander von Humboldt Foundation.

Note: Perform all steps preparatory steps under sterile conditions.
1. Preparation of Plastic Cones for Seed Germination
<ol>
 <li> Fill a 10 cm Petri dish with growth medium containing 1% agar to a thickness of 5 mm. We use a half strength modified Hoagland medium4, but medium composition should be chosen to suit individual experimental requirements. </li>
 <li> While the medium is still liquid, use a multi-channel pipette to fill 10 &mu;l pipette tips with 5 &mu;l of medium from the Petri dish. </li>
 <li> Store the filled tips in the pipette tip box until the medium is solid, then cut to 4 mm long plastic cones and place upright into a Petri dish containing solid growth medium. </li>
</ol>
2. Seed Germination and Seedling Growth 
<ol>
 <li> Surface sterilize seeds in 5% NaOCl for 5 min, wash three times with sterile water, then place a single seed on top of each of the medium-filled cones. </li>
 <li> Seal the dish with Micropore tape (3M) and store at 4 &deg;C to synchronize germination. </li>
 <li> After three days, transfer the plates to a growth cabinet to start germination. Our growth conditions are 23 &deg;C at a 16h high light/8h dark cycle (Light intensity: 100 &mu;E m-2 s-1). </li>
 <li> Between 5 and 7 days after germination, the seedlings should be ready for transfer to the RootChip. At this time, root tips should be near the bottom outlets of the plastic cones. Check seedling health, root length and, if applicable, expression of a fluorescent marker under a dissecting microscope. </li>
 <li> Mark individual seedlings for transfer onto the chip. Select ten or so seedlings in case one is damaged during transfer. </li>
</ol>

3. Transfer of Seedlings onto the RootChip
<ol>
 <li> To sterilize the RootChip for long-term experiments, wrap the device in tissue paper, place in a glass Petri dish, and autoclave. </li>
 <li> Once the RootChip has cooled, cover it with liquid growth medium. The RootChip should be completely immersed, but the fluid level should be no more than 3 mm above the RootChip surface. </li>
 <li> With a 20 &mu;l pipette, pull medium through the root inlet and chamber outlet to fill the observation chamber with medium. </li>
 <li> Plug plastic cones selected in step 2.5 into the RootChip inlets. The cones should fit snugly in the inlets. Since the RootChip is mounted on a thin layer of optical glass, do not apply too much pressure to the chip. </li>
 <li> Incubate the RootChip overnight in liquid medium. To prevent floating, place two glass slides onto the chip. Add a magnetic stir bar and close the dish. </li>
 <li> Transfer the assembly to a magnetic stirrer and gently agitate the medium. </li>
 <li> The RootChip's inlets intersect the channels at a 30&deg; angle to the device normal to facilitate root growth into the channels (Figure 1A). To further support growth in the desired direction, tilt the assembly slightly by placing a glass slide under the Petri dish on the side of the chip opposite of the outlets. </li>
 <li> To maintain the light/ dark cycle, illuminate the seedlings with a ring lamp (Light intensity: 100 &mu;E m-2 s-1) connected to a timer. </li>
</ol>
4. Connecting the RootChip to the Carrier
<ol>
 <li> The following day, fill a sealable, pressurizable vial with liquid growth medium (Figure 1B). </li>
 <li> Invert the chip carrier and place it on a stable surface. Remove the RootChip from the liquid medium and insert it PDMS side down into the bottom aperture of the chip carrier. Orient the chip so that the side containing the control layer inlets is facing the side of the pressure line tubing connectors in the carrier side wall. </li>
 <li> Dry the cover glass on the bottom of the chip by gently blotting with tissue paper. Secure the RootChip to the carrier with tape and right the whole assembly. </li>
 <li>Tubing connectors are made by cutting flexible plastic microbore tubing (TYGON, 0.20" ID x 0.060" OD) into 5 cm long pieces and connecting them to stainless steel microbore tubes (New England Small Tube, 0.025" OD x 0.013" ID x 0.75" long). Fill the tubing connectors with water using a syringe and plug each tubing connector into the corresponding control layer inlet on the chip. The water will later fill the control layer channels and be used to transmit the pressure to the micromechanical valves.</li>
 <li> Plug the opposite ends of the lines into the media/solution vial(s). Apply pressure to the solution vial with a syringe of air. The increased air pressure within the solution vial will force liquid into the lines. </li>
</ol>
5. Mounting the RootChip at the Microscope
<ol>
 <li> Place the carrier onto the microscope stage. To reduce the possibility of the assembly shifting over the course of the experiment due to vibrations in the room, the carrier should fit exactly into the notches of the stage insert. </li>
 <li> The chip valves and the flow of medium through the chip are controlled by air pressure. Two lines with regulators are branched off of a main pressure line - one is used to control the medium flow through the channels, and the other is connected to solenoid air valves that actuate the push-up valves of the control layer. The solenoid valves are operated from the computer via the USB valve controller (developed by Rafael G&oacute;mez-Sj&ouml;berg, Lawrence Berkeley National Lab). Close both pressure regulators before connecting the chip. </li>
 <li> Add a few ml of water to the reservoirs of the carrier to keep the humidity high within the assembly. This step should be repeated over course of longer experiments to keep the plants from drying out. Keep the volume low to minimize the amount of liquid that could be spilled onto the microscope. For long-term experiments, the outflow from the chip outlets can be guided into the reservoirs of the carrier by connecting the chip outlets to the reservoirs with microbore tubing (see step 4.4). Alternatively, the outflow that accumulates on the chip surface can be collected by pipetting. </li>
 <li>Prepare squared sheets of transparent plastic from sheet protectors (C-Line). Fix the transparent plastic to the carrier by double-sided tape to maintain high humidity in the assembly.</li>
 <li> Position the ring light over the chip and maintain the light/dark cycle. The ring light should be switched off prior to the start of any experiment that uses fluorescent markers, as the direct illumination will interfere with image collection. </li>
</ol>
6. Operating the RootChip using the LabView Interface
The RootChip controller interface for the LabView software platform can be downloaded from our website <a href="http://dpb.carnegiescience.edu/technology/rootchip" target="_blank">http://dpb.carnegiescience.edu/technology/rootchip</a>.
<ol>
 <li> The valves on the chip are closed by applying pressure to the control layer, in this case by opening the solenoid air valves. The controller interface allows actuation of the valves by clicking the button below the valve number. Bright green indicates application of pressure and closing of a chip valve (Figure 2B). Activate all three solution-inlet valves in the controller interface before opening the pressure regulators.
 Note: The controller interface features a feedback loop, which allows monitoring of the system's status. This feature may be activated by clicking on the "Readback" button in the controller interface.</li>
 <li> Open the pressure regulator for the control layer and initially set to 15 psi, then open the regulator for the flow layer and initially set to 5 psi. Depending on the desired flow rate, the pressures may be adjusted later. </li>
 <li> Open the inlet valve for the growth medium of choice to flush the chambers with medium. </li>
 <li> Check flow paths under the microscope. Typically, air is trapped in the flow channels and must be removed. Additionally, the channels of the control layer still contain air that must be forced out and replaced by the water from the tubing connectors (dead-end filling). Both tasks are achieved by flushing each of the eight chambers several times (5 psi) until all air is forced from the channels into the PDMS ("degassing"). 
 Note: The controller interface can be programmed to automate experiments. Such routines may also be used to degas the chip. </li>
</ol>
7. Representative Results
The prime purpose of the RootChip is to combine an imaging platform and a perfusion system in a single device with a high level of integration. To demonstrate the manipulation of the microenvironment of roots we flushed the chambers with dark food coloring (1:4 dilution in hydroponic medium) and measured the exchange of fluid within the chambers. At the recommended pressure of 5 psi we measured a full exchange within 10 seconds at a calculated flow rate of approximately 1.5 &mu;l/min (Figure 3).
We also observed root growth of seedlings, in this case grown in the dark and supplied with 10 mM Glucose as an external energy source (Figure 4). Depending on the growth conditions such as light and composition of the medium, plants can be observed in the RootChip for up to three days.
The RootChip has been used to monitor intracellular glucose and galactose levels in roots expressing genetically encoded nanosensors, based on F&ouml;rster Resonance Energy Transfer (FRET)5-7. Roots in the chip were perfused with square pulses of glucose or galactose solution (Figure 5). The intracellular levels of sugars were monitored and are shown here expressed as a ratio of the intensity of the acceptor fluorophore Citrine to the intensity of the donor ECFP. The rise in ratio indicates accumulation of sugar.
<img src="/files/ftp_upload/4290/4290fig1.jpg" alt="Figure 1" /> 
 Figure 1. RootChip principle.
<ol class="ualpha">
 <li> The RootChip features eight observation chambers for the growth and imaging of roots. The seeds first germinate in plastic cones - fabricated from plastic pipette tips - which are filled with solid medium. The root tip grows through the medium and reaches the chamber where a continuous flow of liquid medium keeps the conditions in the chamber constant. Micromechanical valves (red) control the flow. The chip is mounted on an optical cover glass. 
 Drawing not to scale. (Adapted with permission from Grossmann et al., 2011 Plant Cell.)</li>
 <li> Scheme of a pressurizable solution vial with diaphragm (red).</li>
 </ol>
<img src="/files/ftp_upload/4290/4290fig2.jpg" alt="Figure 2" /> 
 Figure 2. Connecting and mounting the RootChip.
<ol class="ualpha">
 <li>Top view of the fully connected RootChip and carrier mounted on an inverted microscope.</li>
 <li> Scheme illustrating the valving system and the controller interface. An example for a valve setting to guide the fluid flow to a single chamber is shown. While valves 4 through 8 act as single valves, valves 0 through 3 act in groups. With this system an individual chamber can be addressed by activating a combination of valves. </li>
</ol>
<img src="/files/ftp_upload/4290/4290fig3.jpg" alt="Figure 3" /> 
 Figure 3. Exchange of solutions in the observation chambers. Visualization of the exchange of fluid in an observation chamber using dye solution. The image is an overlay of bright field and false-colored intensity of the dye signal.
<img src="/files/ftp_upload/4290/4290fig4.jpg" alt="Figure 4" /> 
 Figure 4. On-chip root growth. Observation of a single growing root expressing a fluorescent FRET nanosensor for glucose/galactose over the course of 20h. Time format: hh:mm; scale bar: 100 &mu;m.
<img src="/files/ftp_upload/4290/4290fig5.jpg" alt="Figure 5" /> 
 Figure 5. Measuring intracellular sugar levels using FRET nanosensors.
<ol class="ualpha">
 <li>The amount of sensor is shown as citrine intensity within the root tip (left). The response of the intracellular FRET nanosensor to the application of glucose or galactose solution is shown as ratiometric images. (Adapted with permission from Grossmann et al., 2011 Plant Cell.) Scale bar: 100 &mu;m.</li>
 <li>Tracing FRET ratio changes as a response to three repeated square pulses of glucose (green) and galactose (red).</li>
</ol>

<ol>
	<li>Grossmann, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+RootChip:+An+Integrated+Microfluidic+Chip+for+Plant+Science.">The RootChip: An Integrated Microfluidic Chip for Plant Science.</a> Plant Cell. 23, 4234-4240 (2011).</li><li>Unger, M. A., Chou, H. P., Thorsen, T., Scherer, A., Quake, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monolithic+microfabricated+valves+and+pumps+by+multilayer+soft+lithography.">Monolithic microfabricated valves and pumps by multilayer soft lithography.</a> Science. 288, 113-116 (2000).</li><li>Okumoto, S., Jones, A., Frommer, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+Imaging+with+Fluorescent+Biosensors:+Advanced+Tools+for+Spatiotemporal+Analysis+of+Biodynamics+in+Cells.">Quantitative Imaging with Fluorescent Biosensors: Advanced Tools for Spatiotemporal Analysis of Biodynamics in Cells.</a> Annu. Rev. Plant Biol. , (2012).</li><li>Loqu&#233;, D., Lalonde, S., Looger, L. L., von Wir&#233;n, N., Frommer, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+cytosolic+trans-activation+domain+essential+for+ammonium+uptake.">A cytosolic trans-activation domain essential for ammonium uptake.</a> Nature. 446, 195-198 (2007).</li><li>Okumoto, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+approach+for+monitoring+cellular+metabolites+and+ions+using+genetically+encoded+biosensors.">Imaging approach for monitoring cellular metabolites and ions using genetically encoded biosensors.</a> Curr. Opin. Biotechnol. 21, 45-54 (2010).</li><li>Fehr, M., Frommer, W. B., Lalonde, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualization+of+maltose+uptake+in+living+yeast+cells+by+fluorescent+nanosensors.">Visualization of maltose uptake in living yeast cells by fluorescent nanosensors.</a> Proc. Natl. Acad. Sci. USA. 99, 9846-9851 (2002).</li><li>Takanaga, H., Chaudhuri, B., Frommer, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GLUT1+and+GLUT9+as+major+contributors+to+glucose+influx+in+HepG2+cells+identified+by+a+high+sensitivity+intramolecular+FRET+glucose+sensor.">GLUT1 and GLUT9 as major contributors to glucose influx in HepG2 cells identified by a high sensitivity intramolecular FRET glucose sensor.</a> Biochim. Biophys. Acta. 1778, 1091-1099 (2008).</li><li>Chaudhuri, B., Hörmann, F., Frommer, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+imaging+of+glucose+flux+impedance+using+FRET+sensors+in+wild-type+Arabidopsis+plants.">Dynamic imaging of glucose flux impedance using FRET sensors in wild-type Arabidopsis plants.</a> J. Exp. Bot. 62, 2411-2417 (2011).</li><li>Meier, M., Lucchetta, E. M., Ismagilov, R. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+stimulation+of+the+Arabidopsis+thaliana+root+using+multi-laminar+flow+on+a+microfluidic+chip.">Chemical stimulation of the Arabidopsis thaliana root using multi-laminar flow on a microfluidic chip.</a> Lab Chip. 10, 2147-2153 (2010).</li><li>Parashar, A., Pandey, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plant-in-chip:+Microfluidic+system+for+studying+root+growth+and+pathogenic+interactions+in+Arabidopsis.">Plant-in-chip: Microfluidic system for studying root growth and pathogenic interactions in Arabidopsis.</a> Appl. Phys. Lett. 98, 263703-26 (2011).</li><li>Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+origins+and+the+future+of+microfluidics.">The origins and the future of microfluidics.</a> Nature. 442, 368-373 (2006).</li><li>Lee, J. N., Park, C., Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solvent+Compatibility+of+Poly(dimethylsiloxane)-Based+Microfluidic+Devices.">Solvent Compatibility of Poly(dimethylsiloxane)-Based Microfluidic Devices.</a> Anal. Chem. 75, 6544-6554 (2003).</li></ol>

No conflicts of interest declared.

The main advantages of the RootChip over conventional growth methods are the minimally invasive preparation for microscopy, the ability to reversibly and repeatedly alter the root environment, and the capacity for continuous observation of developmentally competent and physiologically healthy tissue over a period of several days. Previously, seedlings were grown vertically on gelled media and transferred to a perfusion system immediately before the experiment, which allowed only measuring single roots at a time8. Microfluidic tools have been used for Arabidopsis, but on a low integration level9 or without perfusion control10. The RootChip combines a high level of integration with the ability to automate experiments through precise flow guidance. Another advantage of this platform, characteristic of all microfluidic devices11, is that only minimal amounts of liquid are required to supply the root with the necessary nutrients, even for experiments spanning several days. The RootChip is currently designed as a single-use device, but since production costs of chips are low, the small amounts of consumed reagents makes the chip still very cost-effective. 
There are a few critical steps that must be taken to guarantee the health of the seedlings:
The volume in the plastic cones is only 3-4 &mu;l, which will begin to dry when exposed to air. Hence it is critical that the cones are transferred onto the chip quickly and humidity is kept high until the roots have reached the observation chambers, which will supply them with sufficient water. Steps 4.2 to 4.5 should be performed quickly and without interruption to prevent drying of the seedlings.
Steps 3.5 - 3.8 describe the incubation of the chip in liquid media during which the roots grow into the observation chambers. This step may be skipped by mounting the chip into the carrier immediately and starting constant perfusion with growth medium. However, we recommend soaking in growth medium overnight, as it has some advantages: 1) it creates a humid environment so the seedlings are less likely to become desiccated as they grow into the observation chamber; 2) the chip is soaked in liquid, so degassing (step 6.4) will be faster.
It is important to use media with low solute concentrations. More concentrated solutions may precipitate and clog the channels, especially if the chip is used over several days. 
Once the device is connected to the air pressure line, flow of medium is controlled by changing hydraulic pressure in the valves. To guarantee proper closure of the micromechanical valves, it is important to choose a control pressure that is about three times higher than the flow pressure. The flow pressure should not exceed 15 psi as the fluid will be pushed out of the root inlets. Higher pressures may also cause delamination of the chip, which renders the chip unusable.
A limitation of the RootChip is that PDMS is porous and hydrophobic. While the material is practically inert to aqueous solutions, it may absorb organic compounds12. This can interfere with a rapid exchange of solutions as organic compounds may leak from the material even when the supply of this compound has been stopped at the inlet. Due to the porosity, using organic solvents may cause swelling of the PDMS12.
We continue to optimize the RootChip and extend its utility, for example with roots of crop plants. We believe that by improving access to the root for treatments and observation, microfluidic tools like the RootChip will open up new dimensions of root research.

<table border="1">
<tbody>
 <tr>
 <td>Items</td>
 <td>Source</td>
 <td>Information</td>
 </tr>
 <tr>
 <td>Chip carrier, software and other information.</td>
 <td>Carnegie Institution - DPB</td>
 <td>CAD and CNC files for carrier fabrication, controller software and further information are available for download from the website <a href="http://dpb.carnegiescience.edu/technology/rootchip" target="_blank">http://dpb.carnegiescience.edu/technology/rootchip</a> Carriers can also be ordered from this website.</td>
 </tr>
 <tr>
 <td>RootChip</td>
 <td>Stanford Foundry</td>
 <td>Mask designs and fabrication protocols are available upon request. Ready-to-use RootChips can be ordered from <a href="http://www.stanford.edu/group/foundry/" target="_blank">http://www.stanford.edu/group/foundry/</a></td>
 </tr>
 <tr>
 <td>Chip controller</td>
 <td> -home built- </td>
 <td>The automated valve controller system was originally developed by Rafael G&oacute;mez-Sj&ouml;berg , Lawrence Berkeley National Lab. A detailed instruction how to build your own actuated valve controller can be found at <a href="https://sites.google.com/a/lbl.gov/microfluidics-lab/valve-controllers" target="_blank">https://sites.google.com/a/lbl.gov/microfluidics-lab/valve-controllers </a></td>
 </tr>
</tbody>
</table>

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.

Watch this Scientific Journal Video about Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip at JoVE.com

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

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, ...

time-lapse-fluorescence-imaging-arabidopsis-root-growth-with-rapid

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Bioengineering

19.6K Views.  Carnegie Institution for Science. 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.

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

Demonstrating the Uses of the Novel Gravitational Force Spectrometer to Stretch and Measure Fibrous Proteins

The study of macromolecular structure has become critical to the elucidation of molecular mechanisms and function. There are several limited, but important bioinstruments capable of testing the force dependence of structural features in proteins. Scale has been a limiting parameter on how accurately researchers can peer into the nanomechanical world of molecules, such as nucleic acids, enzymes, and motor proteins that perform life-sustaining work. Atomic force microscopy (AFM) is well tuned to determine native structures of fibrous proteins with a distance resolution on par with electron microscopy. However, in AFM force studies, the forces are typically much higher than a single molecule might experience 1, 2. Optical traps (OT) are very good at determining the relative distance between the trapped beads and they can impart very small forces 3. However, they do not yield accurate absolute lengths of the molecules under study. Molecular simulations provide supportive information to such experiments, but are limited in the ability to handle the same large molecular sizes, long time frames, and convince some researchers in the absence of other supporting evidence2, 4.
The gravitational force spectrometer (GFS) fills a critical niche in the arsenal of an investigator by providing a unique combination of abilities. This instrument is capable of generating forces typically with 98% or better accuracy from the femtonewton range to the nanonewton range. The distance measurements currently are capable of resolving the absolute molecular length down to five nanometers, and relative bead pair separation distances with a precision similar to an optical trap. Also, the GFS can determine stretching or uncoiling where the force is near equilibrium, or provide a graded force to juxtapose against any measured structural changes. It is even possible to determine how many amino acid residues are involved in uncoiling events under physiological force loads 2. Unlike in other methods where there is extensive force calibration that must precede any assay, the GFS requires no such force calibration 5. By complementing the strengths of other methods, the GFS will bridge gaps in understanding the nanomechanics of vital proteins and other macromolecules.

This is a step-by step guide showing the purpose, operation, and representative results from the novel gravitational force spectrometer.

The study of macromolecular structure has become critical to the elucidation of molecular mechanisms and function. There are several limited, but ...

Axon Stretch Growth: The Mechanotransduction of Neuronal Growth

During pre-synaptic embryonic development, neuronal processes traverse short distances to reach their targets via growth cone. Over time, neuronal somata are separated from their axon terminals due to skeletal growth of the enlarging organism (Weiss 1941; Gray, Hukkanen et al. 1992). This mechanotransduction induces a secondary mode of neuronal growth capable of accommodating continual elongation of the axon (Bray 1984; Heidemann and Buxbaum 1994; Heidemann, Lamoureux et al. 1995; Pfister, Iwata et al. 2004).
Axon Stretch Growth (ASG) is conceivably a central factor in the maturation of short embryonic processes into the long nerves and white matter tracts characteristic of the adult nervous system. To study ASG in vitro, we engineered bioreactors to apply tension to the short axonal processes of neuronal cultures (Loverde, Ozoka et al. 2011). Here, we detail the methods we use to prepare bioreactors and conduct ASG. First, within each stretching lane of the bioreactor, neurons are plated upon a micro-manipulated towing substrate. Next, neurons regenerate their axonal processes, via growth cone extension, onto a stationary substrate. Finally, stretch growth is performed by towing the plated cell bodies away from the axon terminals adhered to the stationary substrate; recapitulating skeletal growth after growth cone extension.
Previous work has shown that ASG of embryonic rat dorsal root ganglia neurons are capable of unprecedented growth rates up to 10mm/day, reaching lengths of up to 10cm; while concurrently resulting in increased axonal diameters (Smith, Wolf et al. 2001; Pfister, Iwata et al. 2004; Pfister, Bonislawski et al. 2006; Pfister, Iwata et al. 2006; Smith 2009). This is in dramatic contrast to regenerative growth cone extension (in absence of mechanical stimuli) where growth rates average 1mm/day with successful regeneration limited to lengths of less than 3cm (Fu and Gordon 1997; Pfister, Gordon et al. 2011). Accordingly, further study of ASG may help to reveal dysregulated growth mechanisms that limit regeneration in the absence of mechanical stimuli.

A unique tissue engineering method was developed to elongate numerous nerve fibers in culture by recapitulating axon stretch growth; a form of nervous system growth whereby nerves elongate in conjunction with growth of the enlarging body.

During pre-synaptic embryonic development, neuronal processes traverse short distances to reach their targets via growth cone. Over time, neuronal somata ...

Gold Nanostar Synthesis with a Silver Seed Mediated Growth Method

The physical, chemical and optical properties of nano-scale colloids depend on their material composition, size and shape 1-5. There is a great interest in using nano-colloids for photo-thermal ablation, drug delivery and many other biomedical applications 6. Gold is particularly used because of its low toxicity 7-9. A property of metal nano-colloids is that they can have a strong surface plasmon resonance 10. The peak of the surface plasmon resonance mode depends on the structure and composition of the metal nano-colloids. Since the surface plasmon resonance mode is stimulated with light there is a need to have the peak absorbance in the near infrared where biological tissue transmissivity is maximal 11, 12.
We present a method to synthesize star shaped colloidal gold, also known as star shaped nanoparticles 13-15 or nanostars 16. This method is based on a solution containing silver seeds that are used as the nucleating agent for anisotropic growth of gold colloids 17-22. Scanning electron microscopy (SEM) analysis of the resulting gold colloid showed that 70 % of the nanostructures were nanostars. The other 30 % of the particles were amorphous clusters of decahedra and rhomboids. The absorbance peak of the nanostars was detected to be in the near infrared (840 nm). Thus, our method produces gold nanostars suitable for biomedical applications, particularly for photo-thermal ablation.

We synthesized star shaped gold nanostars using a silver seed mediated growth method. The diameter of the nanostars ranges from 200 to 300 nm and the number of tips vary from 7 to 10. The nanoparticles have a broad surface plasmon resonance mode centered in the near infrared.

The physical, chemical and optical properties of nano-scale colloids depend on their material composition, size and shape 1-5. There is a great interest ...

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.

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 ...

Sample Drift Correction Following 4D Confocal Time-lapse Imaging

The generation of four-dimensional (4D) confocal datasets; consisting of 3D image sequences over time; provides an excellent methodology to capture cellular behaviors involved in developmental processes.&#160; The ability to track and follow cell movements is limited by sample movements that occur due to drift of the sample or, in some cases, growth during image acquisition. Tracking cells in datasets affected by drift and/or growth will incorporate these movements into any analysis of cell position. This may result in the apparent movement of static structures within the sample. Therefore prior to cell tracking, any sample drift should be corrected. Using the open source Fiji distribution 1 &#160;of ImageJ 2,3 and the incorporated LOCI tools 4, we developed the Correct 3D drift plug-in to remove erroneous sample movement in confocal datasets. This protocol effectively compensates for sample translation or alterations in focal position by utilizing phase correlation to register each time-point of a four-dimensional confocal datasets while maintaining the ability to visualize and measure cell movements over extended time-lapse experiments.

Time-lapse microscopy allows the visualization of developmental processes. Growth or drift of samples during image acquisition reduces the ability to accurately follow and measure cell movements during development. We describe the use of open source image processing software to correct for three dimensional sample drift over time.

The generation of four-dimensional (4D) confocal datasets; consisting of 3D image sequences over time; provides an excellent methodology to capture ...

3D Orbital Tracking in a Modified Two-photon Microscope: An Application to the Tracking of Intracellular Vesicles

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a modified two-photon microscope1. As opposed to conventional approaches (raster scan or wide field based on a stack of frames), the 3D orbital tracking allows to localize and follow with a high spatial (10 nm accuracy) and temporal resolution (50 Hz frequency response) the 3D displacement of a moving fluorescent particle on length-scales of hundreds of microns2. The method is based on a feedback algorithm that controls the hardware of a two-photon laser scanning microscope in order to perform a circular orbit around the object to be tracked: the feedback mechanism will maintain the fluorescent object in the center by controlling the displacement of the scanning beam3-5. To demonstrate the advantages of this technique, we followed a fast moving organelle, the lysosome, within a living cell6,7. Cells were plated according to standard protocols, and stained using a commercially lysosome dye. We discuss briefly the hardware configuration and in more detail the control software, to perform a 3D orbital tracking experiment inside living cells. We discuss in detail the parameters required in order to control the scanning microscope and enable the motion of the beam in a closed orbit around the particle. We conclude by demonstrating how this method can be effectively used to track the fast motion of a labeled lysosome along microtubules in 3D within a live cell. Lysosomes can move with speeds in the range of 0.4-0.5 &#181;m/sec, typically displaying a directed motion along the microtubule network8.

In this video protocol we track - at high speed and in three dimensions - fluorescently labeled lysosomes within living cells, using the orbital tracking method in a modified two-photon microscope.

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a ...

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

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 ...

Biomimetic Replication of Root Surface Microstructure using Alteration of Soft Lithography

Biomimetics is the use of chemistry and material sciences to mimic biological systems, specifically biological structures, to better humankind. Recently, biomimetic surfaces mimicking the microstructure of leaf surface, were used to study the effects of leaf microstructure on leaf-environment interactions. However, no such tool exists for roots. We developed a tool allowing the synthetic mimicry of the root surface microstructure into an artificial surface. We relied on the soft lithography method, known for leaf surface microstructure replication, using a two-step process. The first step is the more challenging one as it involves the biological tissue. Here, we used a different polymer and curing strategy, relying on the strong, rigid, polyurethane, cured by UV for the root molding. This allowed us to achieve a reliable negative image of the root surface microstructure including the delicate, challenging features such as root hairs. We then used this negative image as a template to achieve the root surface microstructure replication using both the well-established polydimethyl siloxane (PDMS) as well as a cellulose derivative, ethyl cellulose, which represents a closer mimic of the root and which can also be degraded by cellulase enzymes secreted by microorganisms. This newly formed platform can be used to study the microstructural effects of the surface in root-microorganism interactions in a similar manner to what has previously been shown in leaves. Additionally, the system enables us to track the microorganism&#8217;s locations, relative to surface features, and in the future its activity, in the form of cellulase secretion.

Biomimetics has been previously used as a tool to study leaf-microorganism interactions. However, no such tool exists for roots. Here, we develop a protocol to form synthetic surfaces mimicking root surface microstructure for the study of root-environment interactions.

Biomimetics is the use of chemistry and material sciences to mimic biological systems, specifically biological structures, to better humankind. Recently, ...

Blood Flow Imaging with Ultrafast Doppler

The pulsed-Doppler effect is the main technique used in clinical echography to assess blood flow. Applied with conventional focused ultrasound Doppler modes, it has several limits. Firstly, a finely tuned signal filtering operation is needed to distinguish blood flows from surrounding moving tissues. Secondly, the operator must choose between localizing the blood flows or quantifying them. In the last two decades, ultrasound imaging has undergone a paradigm shift with the emergence of ultrafast ultrasound using unfocused waves. In addition to a hundredfold increase in framerate (up to 10000 Hz), this new technique also breaks the conventional quantification/localization trade-off, offering a complete blood flow mapping of the field of view and a simultaneous access to fine velocities measurements at the single-pixel level (down to 50 &#181;m). This data continuity in both spatial and temporal dimensions strongly improves the tissue/blood filtering process, which results in an increase sensitivity to small blood flow velocities (down to 1 mm/s). In this method paper, we aim to introduce the concept of ultrafast Doppler as well as its main parameters. Firstly, we summarize the physical principles of unfocused wave imaging. Then, we present the Doppler signal processing main steps. Particularly, we explain the practical implementation of the critical tissue/blood flow separation algorithms and on the extraction of velocities from these filtered data. This theoretical description is supplemented by in vitro experiences. A tissue phantom embedding a canal with flowing blood-mimicking fluid is imaged with a research programmable ultrasound system. A blood flow image is obtained and the flow characteristics are displayed for several pixels in the canal. Finally, a review of in vivo applications is proposed, showing examples in several organs such as carotids, kidney, thyroid, brain and heart.

This protocol shows how to apply ultrafast ultrasound Doppler imaging to quantify blood flows. After a 1 s long acquisition, the experimenter has access to a movie of the full field of view with axial velocity values for each pixel every &#8776;0.3 ms (depending on the ultrasound time of flight).

The pulsed-Doppler effect is the main technique used in clinical echography to assess blood flow. Applied with conventional focused ultrasound Doppler ...