Name: optical control of living cells electrical activity by conjugated polymers
Uploaded: 2016-01-28T14:00:00
Description: Watch this Scientific Journal Video about Optical Control of Living Cells Electrical Activity by Conjugated Polymers at JoVE.com

The work was supported by EU through project FP7-PEOPLE-212-ITN 316832-OLIMPIA, 
Telethon - Italy (grants GGP12033 and GGP14022), Fondazione Cariplo (grant ID 2013-0738).

1. Preparation of Photoactive Substrates
<ol>
	<li>Prepare a P3HT:PCBM solution (1:1 w/w) in chlorobenzene at a P3HT concentration of 20 g/L. Mix the solution with a magnetic stirrer for at least 4 hr at 60 &#176;C. Consider a volume of 150 &#181;l of solution for each substrate to be prepared.</li>
	<li>Clean ITO-coated glass slides (Rs = 10 &#937;/sq, 18x18 mm2, thickness 170 &#181;m) with consecutive baths of deionized water, acetone and isopropanol in a sonicator (10 min for each step). Dry th...

<ol>
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Critical steps of the reported protocol for in-vitro optical stimulation of cells mainly concern the choice of the light-sensitive polymer, the thermal sterilization parameters, the intensity and the duration of light stimuli. A P3HT:PCBM thin film was selected here, since it guarantees good temporal and electrochemical stability. However, one should notice that not all light-sensitive polymers can offer analogue performances,22 more specifically upon illumination. In addition, in this case selected t...

The possibility to manipulate the cellular activity with a precise spatial and temporal resolution represents a key strategy in neuroscientific research and in the treatment of neurological and psychiatric disorders.1 Traditional methods are based on electrical stimulation of cells using electrodes positioned in proximity or in contact with the targeted system,2 which can be of different complexity (single cell, cellular network, brain slices, in-vivo brain tissues). During the past century, the use of patch-clamp, metal and substrate-integrated electrodes have provided a detailed picture of the physiology and pathophysiology of single neurons and of the functioning mechanisms of neural networks. However, electrical stimulation suffers from important limitations. The first one is related to a generally poor spatial resolution due to the physical dimensions of the electrodes and their fixed geometry, which cannot be readily adapted to complex organized systems like biological tissues. Also, problems related to the electrodes impedance and of cross-talk between stimulation and recording systems may deteriorate the final signal-to-noise ratio of the measurements.3 On the other hand, the use of light for stimulation may help to overcome many limitations of the electrical approach. First of all, it offers unprecedented spatial (&#60; 1 &#181;m) and temporal resolution (&#60; 1 msec), making it possible to target specific cell types or even sub-cell compartments. In addition it is highly non-invasive since it avoids any physical contact with the tissue of interest and disentangles stimulation from recording. Moreover, both light intensity and wavelength can be precisely regulated and thus diverse stimulation protocols can be applied.3,4
However, the vast majority of animal cells do not present any specific sensitivity to light. Several strategies for optical stimulation have thus been proposed, either exploiting light-sensitive molecular mediators nearby or within the cells, or using photoactive device placed externally, close to the cell. The former category refers to endogenous mechanisms like the stimulation via visible or infrared (IR) light, as well as the use of either photoisomerizable/photocleavable compounds or the genetic expression of photosensitive molecular actuators (optogenetics). The latter class includes techniques for exogenous stimulation achieved with the use of inorganic nano/micro-particles or photoconductive silicon substrates.5 Nevertheless, all these systems have bright sides and drawbacks. In particular, endogenous absorption of cells in the visible range is weak and not reliable, and the concomitant generation of reactive oxygen species may be harmful to the cell. In general, IR is used for inducing local thermal heating due to water absorption, but the extinction coefficient of water is small, thus requiring strong infrared light (from tens to hundreds of W/mm2) that is difficult to deliver via standard microscope optics and may pose safety concerns for in-vivo applications. On the other hand, photo-switchable caging compounds have a time limited action and often require UV light that is hard to deliver due to limited tissue penetration. In addition they suffer from diffusion problems of the activated compounds upon photolysis outside the illuminated area. Finally, optogenetic tools have allowed scientists to target specific cellular subpopulation and sub-compartments and are rapidly emerging as one of the key technologies in neuroscientific research. However, inserting an exogenous DNA segment via a viral vector raises important safety issues, especially in view of adoption on human patients.5,6 For these reasons, research on new materials and devices capable of cell optical manipulation is an extremely hot topic.
Recently, a novel approach based on the use of light-sensitive conjugated polymers, able to efficiently transduce an optical stimulus into a modulation of cell electrical activity, has been proposed. The Cell Stimulation by Polymer Photoexcitation (CSPP) technique exploits many key-enabling features typical of organic semiconductors: they are intrinsically sensitive to light in the visible range;7 they are biocompatible, soft and conformable and their mechanical flexibility allows an intimate interface with tissue both in-vitro and in-vivo8-10. Apart from that, they can be easily functionalized to better adapt to the interface with living cells, and to enable specific excitation, probing and sensing capabilities.11,12 Moreover, they support electronic as well as ionic transport, making them ideal for the combination of electronics ad biology.13,14 Interestingly, they can work in photovoltaic mode, avoiding the need to apply an external bias for efficient cell optical stimulation.15 
The reliability of CSPP technique has been previously demonstrated in several systems, including primary neurons,15,16 astrocytes,17 secondary cell lines18 and explanted retinal tissues.16 In this work, all the steps necessary to fabricate a light-sensitive bio-polymer interface19 for optical stimulation of in-vitro systems are described in detail. As a study case, a prototypical organic photovoltaic blend of region-regular poly(3-hexylthiophene) (rr-P3HT), functioning as the electron donor, and phenyl-C61-butyric-acid-methyl ester (PCBM), acting as the electron acceptor is employed. As the biological system, Human Embryonic Kidney (HEK-293) cells are used. An example of a photostimulation protocol with the relative recording of cell activity via electrophysiological measurements is provided.
The described platform is however of general validity, and it can be easily extended to the use of other conjugated polymers (by properly adjusting the solution preparation process and the deposition parameters), different cell types (by properly changing the cell culture protocol, plating procedure and time requested for cell seeding and proliferation) and different stimulation protocols (light wavelength, stimuli frequency and duration, photoexcitation density).

<table border="0" cellpadding="0" cellspacing="0" width="693">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">rr-P3HT</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td style="width:191px;">698989-5G</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">ITO-coated substrates</td>
			<td style="width:120px;">Nano-CS</td>
			<td style="width:191px;">IT10300100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Fibronectin</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td style="width:191px;">F1141</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">chlorobenzene</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td align="right" style="width:191px;">319996</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PCBM</td>
			<td style="width:120px;">Nano-C</td>
			<td style="width:191px;">Nano-CPCBM-BF</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">acetone&#160;</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td align="right" style="width:191px;">270725</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">isopropyl alcohol</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td align="right" style="width:191px;">563935</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">HEK cells</td>
			<td style="width:120px;">LGC standards srl</td>
			<td style="width:191px;">ATCC-CRL-1573</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">HEPES</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td style="width:191px;">H0887</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PBS</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td style="width:191px;">P5244</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">E-MEM</td>
			<td style="width:120px;">LGC standards srl</td>
			<td style="width:191px;">ATCC-30-2003</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">EDTA</td>
			<td style="width:120px;">Sigma Aldrich</td>
			<td style="width:191px;">E8008-</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">FBS</td>
			<td style="width:120px;">LGC standards srl</td>
			<td style="width:191px;">ATCC-30-2020</td>
			<td></td>
		</tr>
	</tbody>
</table>

optical control of living cells electrical activity by conjugated polymers

Hybrid interfaces between organic semiconductors and living tissues represent a new tool for in-vitro and in-vivo applications. In particular, conjugated polymers display several optimal properties as substrates for biological systems, such as good biocompatibility, excellent mechanical properties, cheap and easy processing technology, and possibility of deposition on light, thin and flexible substrates. These materials have been employed for cellular interfaces like neural probes, transistors for excitation and recording of neural activity, biosensors and actuators for drug release. Recent experiments have also demonstrated the possibility to use conjugated polymers for all-optical modulation of the electrical activity of cells. Several in-vitro study cases have been reported, including primary neuronal networks, astrocytes and secondary line cells. Moreover, signal photo-transduction mediated by organic polymers has been shown to restore light sensitivity in degenerated retinas, suggesting that these devices may be used for artificial retinal prosthesis in the future. All in all, light sensitive conjugated polymers represent a new approach for optical modulation of cellular activity.
In this work, all the steps required to fabricate a bio-polymer interface for optical excitation of living cells are described. The function of the active interface is to transduce the light stimulus into a modulation of the cell membrane potential. As a study case, useful for in-vitro studies, a polythiophene thin film is used as the functional, light absorbing layer, and Human Embryonic Kidney (HEK-293) cells are employed as the biological component of the interface. Practical examples of successful control of the cell membrane potential upon stimulation with light pulses of different duration are provided. In particular, it is shown that both depolarizing and hyperpolarizing effects on the cell membrane can be achieved depending on the duration of the light stimulus. The reported protocol is of general validity and can be straightforwardly extended to other biological preparations.

Cells can be easily cultured on P3HT:PCBM substrates, provided that a suitable adhesion layer is deposited (like the fibronectin used in step 3.2 of the described protocol). P3HT:PCBM optical absorption peaks in the green part of the visible spectrum; however other light-sensitive conjugated polymers can be selected, according to the preferred photostimulation wavelengths range (Figure 2). Biocompatibility of these substrates has been demonstrated not only with cell lines...

Watch this Scientific Journal Video about Optical Control of Living Cells Electrical Activity by Conjugated Polymers at JoVE.com

Optical Control of Living Cells Electrical Activity by Conjugated Polymers

A method for stimulation of in-vitro cell cultures electrical activity with visible light, based on the use of organic semiconducting polymers is described.

Hybrid interfaces between organic semiconductors and living tissues represent a new tool for in-vitro and in-vivo applications. In particular, conjugated ...

The overall goal of this method is to control the cell membrane potential upon stimulation by visible light pulses. The phototransduction process is mediated by a light sensitive conjugated polymer. This method provides a new tool in the technologies field seeing as it represents a valuable alternative to existing methods such as optogenetics and thermal stimulation for optical control of living cells activity.The main advantages of this technique are the prompt integrability with any electrophysiologist type, the potentially high special and temporal resolution, they reduce the invasivity. More over, it allows to avoid a gene transfer which is required by optogenetics. Our method is described here for photostimulation of in vitro cell culture.But it might open also interesting opportunities for in vivo applications. Demonstrating the procedure will be Caterina Bossio and Susana Vaquero Morata to post-docs from our labs with biological and physical chemistry backgrounds respectively. To begin, prepare a solution of the light sensitive conjugated polymer by mixing P3HT and PCBM in chlorobenzine at a one to one relative concentration.Mix the solution with a magnetic stirrer for at least four hours at 60 degrees Celsius. Next, clean indium tin oxide-coated glass slides with consecutive 10-minute baths of deionized water, acetone, and isopropanol in a sonicator. Dry the cover slips with a nitrogen gun, and then treat the substrates by a plasma cleaner.Once clean, set the slide on the chuck of a spincoater, ITO-coated side facing upwards. Next, add 150 microliters of the P3HT PCBM solution to the slide and spincoat the sample. After the spincoater is finsihed, remove the sample, and use a cleanroom swab dipped in acetone to clean the back of the substrate.Sterilize the photoactive substrates by placing them into a covered heat resistant container and heating them at 120 degrees Celsius for two hours. Then transfer the samples to a sterile hood and let them cool. From this point on, maintain the sterility of the samples.Place the photoactive substrates into a six well plate. Given the hydrophobicity of the conjugated polymer layer, make a polydimethylsiloxane well to fix the sample, and confine the solutions as described in the accompanying text protocol. Then, place the photoactive substrates into the wells of the plate, and cover the entire surface of each sample with 500 to 750 microliters of a two milligrams per liter solution of fibronectin in PBS.Incubate the samples at 37 degrees Celsius for at least one hour. After incubation, use a glass pipette to remove the fibronectine solution, and then wash the substrates once with two milliliters of PBS. First, add a total volume of 750 to 1, 000 microliters of complete growth medium per well.Next, plate 15, 000 AGK293 cells per square centimeter of photoactive substrate into each well. Then, incubate the cells for 24 to 48 hours. Cells can be easily cultured on the photoactive substrates, provided that a suitable adhesion layer, like fibronectin, is depositited.To begin, prepare both the extracellular and intracellular solutions as described in the accompanying text protocol. Sterilize the solutions by passing them through a 0.2 micron filter. Then, prewarm the extracellular solution by placing it into a water bath at 37 degrees Celsius.Also, fill a one milliliter syringe with the intracellular solution and equip it with a 34-gauge needle. Keep the intracellular solution in contact with ice to avoid degradation of the ATP. Next, take a cell-coated substrate from the incubator and carefully remove the PDMS well.Then remove the growth medium with a glass pipette and rinse the sample slowly with extracellular solution to avoid cell detachment. Add three milliliters of extracellular solution to each well and then transfer the plate to the sample holder of the electrophysiology station. Once in place, set the reference electrode into the well that is to be tested.Next, prepare a fresh glass micropipette for patch clamp using a micropipette puller. Fill half of the pipette with intracellular solution. Use the micromanipulator to position the patch pipette in close proximity to the selected cell membrane.During the positioning, apply over pressure to the pipette in order to avoid dirt sticking on the tip. With the pipette a few microns above the cell, start lowering the pipette towards the cell membrane while contolling the pipette resistance on the patch control software. When the pipette resistance increases about one megaohm, remove the over pressure and apply a gentle suction in order to form the gigaseal between the pipette and the cell membrane.Next, apply a negative potential of negative 40 millivolts to the pipette to help stabilize the seal. Compensate for the capacitive transience due to the pipette capacitance with the relative commands on the patch amplifier and/or the patch control software. Apply a brief and intense suction impulse to the pipette in order to break the membrane and to allow electrical access to the cell cytoplasm.Then, set the patch amplifier to current clamp mode so that no current is injected into the cell. In current clamp mode, track the cell membrane potential and wait a few minutes for it to stabilize. Once stable, apply the desired illumination protocol to the cell by illuminating the photoactive membrane under the cell with light in the range of 450 to 600 nanometers.Record the effect of the light on the membrane potential. Short pulses of light around 10 to 20 milliseconds result in a transient depolarization of the cell, while with prolonged illumination in order of hundreds of milliseconds are sustained, hyperbolization is observed until the light is switched off. Optical stimulation of cells mediated by the photoactive substrates can result in different effects on the cell membrane, depending on the duration of the light stimulus.A fast spike, like the one shown here, is followed by a transient depolarization on the order of tens of milliseconds. For short pulses of light as the light is switched off, an opposite behavior is observed. These signals have been attributed to a variation in membrane capacitance due to local heating following light absorption.For prolonged illumination, the initial depolarization turns into a cell hyperpolarization. This phenomenon has again a thermal origin, but in this case, it is related to a variation of the membrane equilibrium potential due to a change in the ion channel's conductivities induced by the increased temperature. It is important to note that this technique is quite flexible and can be adapted to the particular needs of a specific experiment, for instance, by changing the photoxic polymer, the fabrication process, or the illumination protocol.However, before changing the protocol, one should first verify the important properties of the active polymer like its biocompatability, suitability to the sterilization process, electrochemical and temporal stability. After watching this video, you should have a good understanding of how to realize a hybrid interface between connective polymers and living cells and how to use it for optical control of cell activity. We believe that this technique also promises to become a complimentary tool in neuroscience for in vitro investigations and also to open a new perspective in vivo applications.To refine the protocol, however, a strong and joint effort from material scientists and neuroscientists, physicists, and medical doctors is mandatory.

optical-control-living-cells-electrical-activity-conjugated

Research

JoVE Journal

Bioengineering

Shape Memory Polymers for Active Cell Culture

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent shape upon the application of a stimulus such as heat1-5. In a typical shape memory cycle, the SMP is first deformed at an elevated temperature that is higher than its transition temperature, Ttrans [either the melting temperature (Tm) or the glass transition temperature (Tg)]. The deformation is elastic in nature and mainly leads to a reduction in conformational entropy of the constituent network chains (following the rubber elasticity theory). The deformed SMP is then cooled to a temperature below its Ttrans while maintaining the external strain or stress constant. During cooling, the material transitions to a more rigid state (semi-crystalline or glassy), which kinetically traps or "freezes" the material in this low-entropy state leading to macroscopic shape fixing. Shape recovery is triggered by continuously heating the material through Ttrans under a stress-free (unconstrained) condition. By allowing the network chains (with regained mobility) to relax to their thermodynamically favored, maximal-entropy state, the material changes from the temporary shape to the permanent shape.
Cells are capable of surveying the mechanical properties of their surrounding environment6. The mechanisms through which mechanical interactions between cells and their physical environment control cell behavior are areas of active research. Substrates of defined topography have emerged as powerful tools in the investigation of these mechanisms. Mesoscale, microscale, and nanoscale patterns of substrate topography have been shown to direct cell alignment, cell adhesion, and cell traction forces7-14. These findings have underscored the potential for substrate topography to control and assay the mechanical interactions between cells and their physical environment during cell culture, but the substrates used to date have generally been passive and could not be programmed to change significantly during culture. This physical stasis has limited the potential of topographic substrates to control cells in culture.
Here, active cell culture (ACC) SMP substrates are introduced that employ surface shape memory to provide programmed control of substrate topography and deformation. These substrates demonstrate the ability to transition from a temporary grooved topography to a second, nearly flat memorized topography. This change in topography can be used to control cell behavior under standard cell culture conditions.

A method for developing cell culture substrates with the ability to change topography during culture is described. The method makes use of smart materials known as shape memory polymers that have the ability to memorize a permanent shape. This concept is adaptable to a wide range of materials and applications.

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent ...

Fluorescence Lifetime Imaging of Molecular Rotors in Living Cells

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular events. Viscosity is one of the key parameters affecting the diffusion of molecules and proteins, and changes in viscosity have been linked to disease and malfunction at the cellular level.1-3 While methods to measure the bulk viscosity are well developed, imaging microviscosity remains a challenge. Viscosity maps of microscopic objects, such as single cells, have until recently been hard to obtain. Mapping viscosity with fluorescence techniques is advantageous because, similar to other optical techniques, it is minimally invasive, non-destructive and can be applied to living cells and tissues.
Fluorescent molecular rotors exhibit fluorescence lifetimes and quantum yields which are a function of the viscosity of their microenvironment.4,5 Intramolecular twisting or rotation leads to non-radiative decay from the excited state back to the ground state. A viscous environment slows this rotation or twisting, restricting access to this non-radiative decay pathway. This leads to an increase in the fluorescence quantum yield and the fluorescence lifetime. Fluorescence Lifetime Imaging (FLIM) of modified hydrophobic BODIPY dyes that act as fluorescent molecular rotors show that the fluorescence lifetime of these probes is a function of the microviscosity of their environment.6-8 A logarithmic plot of the fluorescence lifetime versus the solvent viscosity yields a straight line that obeys the F&ouml;rster Hoffman equation.9 This plot also serves as a calibration graph to convert fluorescence lifetime into viscosity.
Following incubation of living cells with the modified BODIPY fluorescent molecular rotor, a punctate dye distribution is observed in the fluorescence images. The viscosity value obtained in the puncta in live cells is around 100 times higher than that of water and of cellular cytoplasm.6,7 Time-resolved fluorescence anisotropy measurements yield rotational correlation times in agreement with these large microviscosity values. Mapping the fluorescence lifetime is independent of the fluorescence intensity, and thus allows the separation of probe concentration and viscosity effects. 
In summary, we have developed a practical and versatile approach to map the microviscosity in cells based on FLIM of fluorescent molecular rotors.

Fluorescence Lifetime Imaging (FLIM) has emerged as a key technique to image the environment and interaction of specific proteins and dyes in living cells. FLIM of fluorescent molecular rotors allows mapping of viscosity in living cells.

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular ...

Spatio-Temporal Manipulation of Small GTPase Activity at Subcellular Level and on Timescale of Seconds in Living Cells

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular functions and events1, 2. Their spatiotemporally dynamic nature has been revealed by visualization of their activity and localization in real time3. In order to gain deeper understanding of their roles in diverse cellular functions at the molecular level, the next step should be perturbation of protein activities at a precise subcellular location and timing. 
To achieve this goal, we have developed a method for light-induced, spatio-temporally controlled activation of small GTPases by combining two techniques: (1) rapamycin-induced FKBP-FRB heterodimerization and (2) a photo-caging method of rapamycin. With the use of rapamycin-mediated FKBP-FRB heterodimerization, we have developed a method for rapidly inducible activation or inactivation of small GTPases including Rac4, Cdc424, RhoA4 and Ras5, in which rapamycin induces translocation of FKBP-fused GTPases, or their activators, to the plasma membrane where FRB is anchored. For coupling with this heterodimerization system, we have also developed a photo-caging system of rapamycin analogs. A photo-caged compound is a small molecule whose activity is suppressed with a photocleavable protecting group known as a caging group. To suppress heterodimerization activity completely, we designed a caged rapamycin that is tethered to a macromolecule such that the resulting large complex cannot cross the plasma membrane, leading to virtually no background activity as a chemical dimerizer inside cells6. Figure 1 illustrates a scheme of our system. With the combination of these two systems, we locally recruited a Rac activator to the plasma membrane on a timescale of seconds and achieved light-induced Rac activation at the subcellular level6.

A method for spatio-temporal control of small GTPase activity by light is described. This method is based on rapamycin-induced FKBP-FRB heterodimerization and photo-caging systems. Optimization of light-irradiation enables the spatio-temporally controlled activation of small GTPases at the subcellular level.

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular ...

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Optical Detection of E. coli Bacteria by Mesoporous Silicon Biosensors

A label-free optical biosensor based on a nanostructured porous Si is designed for rapid capture and detection of Escherichia coli K12 bacteria, as a model microorganism. The biosensor relies on direct binding of the target bacteria cells onto its surface, while no pretreatment (e.g.&#160;by cell lysis) of the studied sample is required. A mesoporous Si thin film is used as the optical transducer element of the biosensor. Under white light illumination, the porous layer displays well-resolved Fabry-P&#233;rot fringe patterns in its reflectivity spectrum. Applying a fast Fourier transform (FFT) to reflectivity data results in a single peak. Changes in the intensity of the FFT peak are monitored. Thus, target bacteria capture onto the biosensor surface, through antibody-antigen interactions, induces measurable changes in the intensity of the FFT peaks, allowing for a &#39;real time&#39; observation of bacteria attachment.
The mesoporous Si film, fabricated by an electrochemical anodization process, is conjugated with monoclonal antibodies, specific to the target bacteria. The immobilization, immunoactivity and specificity of the antibodies are confirmed by fluorescent labeling experiments. Once the biosensor is exposed to the target bacteria, the cells are directly captured onto the antibody-modified porous Si surface. These specific capturing events result in intensity changes in the thin-film optical interference spectrum of the biosensor. We demonstrate that these biosensors can detect relatively low bacteria concentrations (detection limit of 104 cells/ml) in less than an hour.

Optical Detection of E. coli Bacteria by Mesoporous Silicon Biosensors

A label-free optical biosensor for rapid bacteria detection is introduced. The biosensor is based on a nanostructured porous Si, which is designed to directly capture the target bacteria cells onto its surface. We use monoclonal antibodies, immobilized onto the porous transducer, as the capture probes. Our studies demonstrate the applicability of these biosensors for the detection of low bacterial concentrations within minutes with no prior sample processing (such as cell lysis).

A label-free optical biosensor based on a nanostructured porous Si is designed for rapid capture and detection of Escherichia coli K12 bacteria, as a ...

Nanomanipulation of Single RNA Molecules by Optical Tweezers

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be increasingly important. As RNA function often depends on its ability to adopt alternative structures, it is difficult to predict RNA three-dimensional structures directly from sequence. Single-molecule approaches show potentials to solve the problem of RNA structural polymorphism by monitoring molecular structures one molecule at a time. This work presents a method to precisely manipulate the folding and structure of single RNA molecules using optical tweezers. First, methods to synthesize molecules suitable for single-molecule mechanical work are described. Next, various calibration procedures to ensure the proper operations of the optical tweezers are discussed. Next, various experiments are explained. To demonstrate the utility of the technique, results of mechanically unfolding RNA hairpins and a single RNA kissing complex are used as evidence. In these examples, the nanomanipulation technique was used to study folding of each structural domain, including secondary and tertiary, independently. Lastly, the limitations and future applications of the method are discussed.

Optical tweezers have been used to study RNA folding by stretching individual molecules from their 5&#8217; and 3&#8217; ends. Here common procedures are described to synthesize RNA molecules for tweezing, calibration of the instrument, and methods to manipulate single molecules.

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be ...

Transmission of Multiple Signals through an Optical Fiber Using Wavefront Shaping

The transmission of multiple independent optical signals through a multimode fiber is accomplished using wavefront shaping in order to compensate for the light distortion during the propagation within the fiber. Our methodology is based on digital optical phase conjugation employing only a single spatial light modulator, where the optical wavefront is individually modulated at different regions of the modulator, one region per light signal. Digital optical phase conjugation approaches are considered to be faster than other wavefront shaping approaches, where (for example) a complete determination of the wave propagation behavior of the fiber is performed. In contrast, the presented approach is time-efficient since it only requires one calibration per light signal. The proposed method is potentially appropriate for spatial division multiplexing in communications engineering. Further application fields are endoscopic light delivery in biophotonics, especially in optogenetics, where single cells in biological tissue have to be selectively illuminated with high spatial and temporal resolution.

We demonstrate the transmission of multiple independent signals through a multimode fiber using wavefront shaping employing a single spatial light modulator. By modulating the wavefront for each signal individually, spatially separated foci are transmitted. Potential applications are multiplexed data transfer in communications engineering and endoscopic light delivery in biophotonics.

The transmission of multiple independent optical signals through a multimode fiber is accomplished using wavefront shaping in order to compensate for the ...

Using Extraordinary Optical Transmission to Quantify Cardiac Biomarkers in Human Serum

For a biosensing platform to have clinical relevance in point-of-care (POC) settings, assay sensitivity, reproducibility, and ability to reliably monitor analytes against the background of human serum are crucial.
Nanoimprinting lithography (NIL) was used to fabricate, at a low cost, sensing areas as large as 1.5 mm x 1.5 mm. The sensing surface was made of high-fidelity arrays of nanoholes, each with an area of about 140 nm2. The great reproducibility of NIL made it possible to employ a one-chip, one-measurement strategy on 12 individually manufactured surfaces, with minimal chip-to-chip variation. These nanoimprinted localized surface plasmon resonance (LSPR) chips were extensively tested on their ability to reliably measure a bioanalyte at concentrations varying from 2.5 to 75 ng/mL amidst the background of a complex biofluid-in this case, human serum. The high fidelity of NIL enables the generation of large sensing areas, which in turn eliminates the need for a microscope, as this biosensor can be easily interfaced with a commonly available laboratory light source. These biosensors can detect cardiac troponin in serum with a high sensitivity, at a limit of detection (LOD) of 0.55 ng/mL, which is clinically relevant. They also show low chip-to-chip variance (due to the high quality of the fabrication process). The results are commensurable with widely used enzyme-linked immunosorbent assay (ELISA)-based assays, but the technique retains the advantages of an LSPR-based sensing platform (i.e., amenability to miniaturization and multiplexing, making it more feasible for POC applications).

This work describes a nanoimprinting lithography method to fabricate high-quality sensing arrays that work on the principle of extraordinary optical transmission. The biosensor is low-cost, robust, easy to use, and can detect cardiac troponin I in serum at clinically relevant concentrations (99th percentile cutoff &#8764;10-400 pg/mL, depending on the assay).

For a biosensing platform to have clinical relevance in point-of-care (POC) settings, assay sensitivity, reproducibility, and ability to reliably monitor ...

Trapping of Micro Particles in Nanoplasmonic Optical Lattice

The plasmonic optical tweezer has been developed to overcome the diffraction limits of the conventional far field optical tweezer. Plasmonic optical lattice consists of an array of nanostructures, which exhibit a variety of trapping and transport behaviors. We report the experimental procedures to trap micro-particles in a simple square nanoplasmonic optical lattice. We also describe the optical setup and the nanofabrication of a nanoplasmonic array. The optical potential is created by illuminating an array of gold nanodiscs with a Gaussian beam of 980 nm wavelength, and exciting plasmon resonance. The motion of particles is monitored by fluorescence imaging. A scheme to suppress photothermal convection is also described to increase usable optical power for optimal trapping. Suppression of convection is achieved by cooling the sample to a low temperature, and utilizing the near-zero thermal expansion coefficient of a water medium. Both single particle transport and multiple particle trapping are reported here.

We describe a procedure to optically trap micro-particles in nanoplasmonic optical lattice.

The plasmonic optical tweezer has been developed to overcome the diffraction limits of the conventional far field optical tweezer. Plasmonic optical ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...