Name: bridging the bio-electronic interface with biofabrication
Uploaded: 2012-06-06T13:00:00
Duration: 16 min 38 s
Description: Watch this Scientific Journal Video about Bridging the Bio-Electronic Interface with Biofabrication at JoVE.com

We acknowledge funding from DTRA for support of this manuscript and from ONR, DTRA, and NSF for partial support of underlying research.

1. Alginate Electrodeposition
<ol>
 <li>Connect a power supply to the custom fabricated electrodes via patch cords using alligator clips. An indium tin oxide (ITO) covered glass slide will act as the anode (working electrode) and platinum foil will serve as the cathode (counter-electrode). Position the electrodes so the ITO surface to be functionalized opposes the counter electrode and is positioned either vertically to be dipped into a solution or horizontally such that the deposition solution is contained on the surface.</li>
 <li>Prepare an alginate deposition solution by mixing 1% alginate and 0.5% CaCO3 (by weight) in distilled water and then autoclaving the solution. It is recommended to continuously stir the solution when not in use.</li>
 <li>Submerge both electrodes into the deposition solution. The alginate used may be fluorescently labeled with FluoroSpheres (Invitrogen), as per Cheng et al. 14, to allow for fluorescence imaging of the resulting film.</li>
 <li>Apply a constant current density (3 A/m2) for 2 minutes; voltage will shift within the range of 2-3 V.</li>
 <li>Disconnect the electrode and remove the non-deposited solution. Gently rinse the film with NaCl (0.145 M) to remove superfluous alginate.</li>
 <li>Incubate the film briefly (~1 min) in CaCl2 (0.1 M) to strengthen the gel. Rinse with NaCl (0.145 M) and incubate in a desired solution supplemented with CaCl2 (1 mM).</li>
 <li>Image using a fluorescence microscope (Figure 1B).</li>
</ol>
2. Codeposition of Communicating Cell Populations in Alginate
<ol>
 <li>A signal-sender cell culture (W3110 wt E. coli), grown in LB media, and a signal-receiver cell culture (MDAI2 + pCT6-lsrR-ampr + pET-dsRed-kanr), grown in LB media + 50 &mu;g/mL each of kanamycin and ampicillin, must be grown up overnight and re-inoculated for growth to an optical density (at 600 nm) of 1. Adjust optical density of the receiver cell culture to 0.4-0.6 with LB before use.</li>
 <li>Prepare a deposition solution of 2% alginate and 1% CaCO3, and mix with each cell culture in a 1:1 ratio to a final concentration of 1% alginate, 0.5% CaCO3, with the cells diluted to a density of about half the culturing density.</li>
 <li>Use a glass slide patterned with two ITO electrodes contained by a polydimethylsiloxane (PDMS) well (prepared according to Sylgard instructions and cut to desired size) and a platinum counter electrode. Connect one ITO electrode to a power supply with the platinum as described in Procedure 1 (Alginate electrodeposition).</li>
 <li>Immerse the electrodes in a deposition solution containing the receiver cells. Set the power supply to a constant current at a density of 3 A/m2, where the surface area dimension is defined by the single electrode on which deposition will occur.</li>
 <li>Apply the current for 2 minutes to allow for codeposition of the cells in the alginate matrix.</li>
 <li>Rinse the film as described in Step 1.5.</li>
 <li>Switch the anodic connection to the second, adjacent, ITO electrode.</li>
 <li>Repeat the deposition procedure (Steps 2.4 - 2.7), but this time introducing the solution containing the sender cells.</li>
 <li>Incubate the 2-electrode chip containing codeposited cells and calcium-alginate overnight at 37 &deg;C in phosphate buffered saline (PBS) supplemented with 10% LB media and 1 mM CaCl2.</li>
 <li>After incubation, image using a fluorescence microscope (Figure 2B).</li>
</ol>
3. Chitosan Electrodeposition
<ol>
 <li>Connect the power supply to the electrodes via alligator clips. A gold coated silicon chip will act as the cathode (working electrode) and a platinum foil will serve as the anode (counter-electrode). Position the gold electrode's surface so that it opposes the counter electrode and both are positioned either vertically to be dipped into a solution or horizontally such that the deposition solution is contained on the surface.</li>
 <li>Prepare a chitosan solution by mixing chitosan flakes into water and slowly adding 2 M HCl to dissolve the polysaccharides (final pH 5.6), making sure to follow the procedure outlined by Meyer et al. 15.</li>
 <li>Place the electrodes into a chitosan solution (0.8 %), completely submerging the desired area for deposition. The chitosan used can be fluorescently labeled with 5-(and-6)-carboxyrhodamine 6G succinimidyl ester (Invitrogen), as per Wu et al. 8, to image the electrodeposited film by fluorescence microscopy.</li>
 <li>Apply a constant current density (4 A/m2) for 2 minutes. Voltage will shift within the range of 2-3 V. Calculate the current density as a function of the gold surface area of the working electrode exposed to the deposition solution.</li>
 <li>Rinse the electrode with DI water to remove superfluous chitosan. The chip can be stored in water or PBS (10 mM, pH 7.0).</li>
 <li>Image using a fluorescence microscope (Figure 3C).</li>
</ol>
4. Electrochemical Transduction with a Functionalized Chitosan Film
<ol>
 <li>Codeposit chitosan and glucose oxidase (GOx) from a solution (1% chitosan, 680 U/mL GOx, pH 5.6) at a current density of 4 A/m2 onto a patterned electrode according to Procedure 3 (Chitosan electrodeposition). A chitosan film entrapped in GOx will be generated.</li>
 <li>Attach the treated electrode to a three-electrode system as the working electrode, a platinum wire as the counter-electrode and Ag/AgCl as the reference electrode, as described in Figure 4A.</li>
 <li>Immerse the electrodes into a phosphate buffer solution (0.1 M, pH 7.0) containing NaCl (0.1 M).</li>
 <li>Electrochemically conjugate the protein to the chitosan film by applying a constant voltage (0.9 V) for 60s using chronoamperometry10.</li>
 <li>Place the chip in phosphate buffer (0.1 M, pH 7.0) and wash for 10 minutes on an orbital shaker to remove any unreacted NaCl and unconjugated GOx.</li>
 <li>Re-attach to the three-electrode system as described in step 4.2 and immerse into a solution of 5 mM glucose. Using cyclic voltammetry, sweep the potential in a positive direction to 0.7 V. Use a control film containing no glucose oxidase as a comparison for the amount of oxidation seen in the sweep (Figure 4B).</li>
 <li>Remove the electrodes from the glucose solution and rinse with phosphate buffer (0.1 M, pH 7.0) then place the electrodes into a 10 mL beaker containing 8 mL of phosphate buffer (0.1 M, pH 7.0). Bias the GOx-functionalized chip to 0.6 V to serve as the working electrode (Figure 4C).</li>
 <li>Add aliquots of glucose to the buffer (each aliquot increases the glucose concentration by 4 mM).</li>
 <li>Generate a standard curve between the steady state current and the glucose concentration for the GOx-functionalized chitosan film.</li>
</ol>
5. Protein Functionalization Using Enzymatic Assembly
<ol>
 <li>Use a glass slide patterned with an adjacent gold and ITO electrode contained within a PDMS well. Bias the gold electrode with a cathodic potential to electrodeposit chitosan as shown previously. Rinse film briefly in DI water and then PBS by pipette.</li>
 <li>Add a solution of 3 &mu;M blue fluorescently labeled "AI-2 Synthase" 16 (using a DyLight labeling kit) + 100 U/mL Tyrosinase in PBS.</li>
 <li>Incubate for 1 h at room temperature, then rinse the film with PBS.</li>
 <li>Apply an anodic potential to the ITO electrode to codeposit an alginate deposition solution containing receiver cells (as prepared in steps 2.1-2.2). Follow steps 2.3-2.6 of Procedure 2 (Codeposition of cell populations in alginate).</li>
 <li>To generate the transmitted signal (AI-2) enzymatically, after rinsing the films add a solution of 500 &mu;M S-adenosyl homocysteine (SAH) in PBS, supplemented with 10% LB media and 1mM CaCl2. Cover the electrodes to prevent evaporation of the solution and incubate overnight at 37 &deg;C. This will allow for a receiver cell response by generating a red fluorescent protein (dsRed).</li>
 <li>Adjacent electrodes may be imaged with fluorescence microscopy by adjusting the filters to capture the blue fluorescence of the AI-2 Synthase and red fluorescence expressed by the codeposited receiver cells (Figure 5B).</li>
</ol>
6. Representative Results
Imposed electrical signals can create localized microenvironments (e.g., fields and gradients) near an electrode surface and these stimuli can trigger the self-assembly of polysaccharides such as alginate and chitosan to deposit as a hydrogel film on the electrode surface. Because this sol-gel transition occurs at the electrode surface, the resulting film is electroaddressed, with its geometry matching the electrode pattern (Figures 1B, 3C). Biocompatible films such as alginate and chitosan provide surfaces that can be functionalized with biological components. Using alginate, unique cell populations have been codeposited at separate addresses. Evidence of their electroaddressment is observed upon interaction between the sender and receiver cell population. The molecule autoinducer-2 (AI-2) diffuses from the sender cells and is taken up by the receiver cells, resulting in expression of the dsRed red fluorescent protein (Figure 2A). In Figure 2B, red fluorescence is observed only at the electrode where receivers are addressed.
The amine groups present on chitosan provide it with the pH responsiveness required for electrodeposition as well as a surface suitable for functionalization. We utilized these unique properties by electrochemically conjugating the biosensing enzyme glucose oxidase (GOx) to electrodeposited chitosan films. This enzyme then provides the ability for detection of glucose through an enzymatic reaction (Figure 4A) producing hydrogen peroxide which can then be electrochemically oxidized to produce an output current. In this way, a chemical signal can be transduced to electrical. Figure 4B shows that films in which GOx was electrochemically conjugated produce a strong anodic signal in the presence of glucose as opposed to those films containing no GOx. These results indicate GOx can be assembled onto a deposited chitosan film and will retain catalytic activity. Furthermore, Figure 4C shows a step-increase in anodic current produced in response to increasing glucose concentrations. The standard curve also present in Figure 4C shows that the step-increases proceeded in a near linear fashion dependent on the amount of glucose added. These results show that the enzyme also retains its sensitivity to increasing glucose concentrations upon conjugation to the chitosan film. The lower limit of detection was not studied here as it has been previously characterized for this system in the work of Meyer et. al.
We also have demonstrated the covalent immobilization of an enzyme of interest, engineered to contain a custom penta-tyrosine tag, to chitosan in an enzymatically-controlled manner. Specifically, this process is mediated by the enzyme tyrosinase. As depicted by the scheme in Figure 5A(upper), an enzyme, AI-2 Synthase includes a penta-tyrosine tag. Tyrosinase acts on the tyrosine tag, oxidizing the residues' phenol groups to O-quinones, which then covalently bind to chitosan's amines. Evidence of chitosan film functionalization with the AI-2 Synthase by tyrosinase assembly is observed in Figure 5B, where the AI-2 Synthase has been fluorescently labeled blue. Because AI-2 Synthase generates AI-2 from the substrate S-adenosyl homocysteine (SAH) in the same way as the sender cells, its proximity to codeposited receiver cells in the presence of SAH also causes the receiver cells to fluorescently respond by expressing dsRed (Figure 5A(lower)). Red fluorescence of the receiver cells (Figure 5B) again demonstrates interaction between addresses due to the diffusion of AI-2 from one to the other, and further indicates that enzymes immobilized to chitosan retain activity once covalently bound.
<img alt="Figure 1" src="/files/ftp_upload/4231/4231fig1.jpg" /> 
Figure 1. Alginate electrodeposition. (A) Mechanism of alginate electrodeposition: As an electrode is anodically biased, water electrolysis occurs at its surface, generating a localized low pH. Calcium carbonate particles react with the surplus of protons, releasing calcium cations as the particles dissolve. In the presence of alginate polymer chains, the ions become chelated in an "eggbox" network, forming a crosslinked hydrogel at the electrode. As the distance from the electrode increases, alginate has a greater tendency to remain in solution due to the reduced presence of calcium ions. (B) An L-shaped patterned ITO electrode was used to electrodeposit alginate. A PDMS well was fixed to the electrode to contain a green fluorescently-labeled alginate (1%) and CaCO3 (0.5%) deposition solution. After electrodepositing for 2 min. at a current density of 3A/m2, the electroaddressed alginate hydrogel was imaged by fluorescence microscopy.
<img alt="Figure 2" src="/files/ftp_upload/4231/4231fig2.jpg" /> 
Figure 2. Codeposition of cell populations. (A) Scheme showing interaction between two E. coli strains: One population produces autoinducer-2 (AI-2), a signaling molecule, and is termed "AI-2 sender." The other population, termed "AI-2 receiver," is a reporter of AI-2; upon receipt of AI-2 by diffusion from the sender, it expresses the red fluorescent protein dsRed. (b) Red fluorescence image of electrode pair with the AI-2 sender population codeposited with alginate on the left electrode and AI-2 receiver population codeposited with alginate on right electrode. Magnified view demonstrates the dsRed expression of only the AI-2 receivers.
<img alt="Figure 3" src="/files/ftp_upload/4231/4231fig3.jpg" /> 
Figure 3. Chitosan electrodeposition. (A) Scheme showing the pH-dependent electrodeposition of chitosan. Water electrolysis at a cathodically biased electrode causes a localized high pH (shown by a localized color change of a pH indicator dye near the cathode in micrograph) which stimulates the sol-gel transition of chitosan in this region. (B) The amines present on chitosan give it pH-responsive properties. Above a pH of 6.3 (pKa of chitosan) the amines are deprotonated, facilitating a transition from its protonated soluble form to its insoluble gel form. (C) A patterned gold electrode was used to electrodeposit chitosan. The electrode, connected cathodically to the power supply, was immersed into a green fluorescently-labeled chitosan (0.8%) deposition solution. After electrodepositing for 2 min. at a current density of 4 A/m2, the electroaddressed chitosan film was imaged by fluorescence microscopy.
<img alt="Figure 4" src="/files/ftp_upload/4231/4231fig4.jpg" /> 
Figure 4. Electrochemical transduction with a functionalized chitosan film. (A) Schematic showing the set-up of a three electrode system. Functionalized chitosan film serves as the working electrode, a platinum wire as the counter electrode and Ag/AgCl as the reference electrode. Electrochemical transduction of glucose proceeds through the enzymatic and electrochemical reactions shown where produced hydrogen peroxide can be oxidized and detected at the working electrode. (B) Cyclic voltammagram (CV) at electrode with a chitosan film containing electrochemically conjugated glucose oxidase (GOx) shows a strong anodic signal in a 5 mM glucose solution. A film containing no GOx served as a control and displayed no signal in the same solution. (C) A standard curve between anodic current and the glucose concentration displays a near linear relationship (each aliquot increased the glucose concentration by 4 mM and also increased the current amplitude in the inset graph in a step-wise manner).
<img alt="Figure 5" src="/files/ftp_upload/4231/4231fig5.jpg" /> 
Figure 5. Protein functionalization using enzymatic assembly. (A, upper) Scheme showing tyrosine-tagged "AI-2 Synthase" being covalently tethered to a chitosan film by tyrosinase assembly. The tyrosine residues become oxidized to O-quinones by tyrosinase action and may react with amine groups on the chitosan film, forming a covalent bond. (A, lower) The AI-2 Synthase generates AI-2 from a substrate (SAH); the receiver cells report the generated AI-2 by dsRed fluorescence expression. (B) Fluorescence images showing a chitosan film on gold, functionalized with blue-labeled AI-2 Synthase. Adjacently, AI-2 receiver cells are codeposited with alginate on ITO. After addition of the enzymatic substrate to the well and incubation, the AI-2 receiver cells express dsRed.

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Our procedures demonstrate the electrodeposition and functionalization of biopolymer films, a process we term biofabrication. Through functionalization with cells and biomolecules we create biological surfaces capable of interacting with each other and the electrode address they are assembled upon. The first step, electrodeposition, takes place through the triggered self-assembly of biopolymers, alginate and chitosan in our studies, in response to an electrical signal. As stated earlier a pH gradient is generated which can be controlled by the current density and deposition time, providing additional control over the film dimensions and properties 6,17. We have found that a variety of current density and deposition time combinations can be used for the electrodes indicated in Table 1. While use of other electrodes is feasible, adjustments to the procedure would be necessary. Compared with other techniques of film formation the process of electrodeposition is simple, rapid and reagentless. There is no need for an extensive repertoire of expensive equipment and laborious preparations. Importantly, the process can withstand minor experimental deviations and can be easily started over if a problem occurs. 
Chitosan is capable of responding to a high cathodic pH gradient due to important functional properties conferred to it by a high content of primary amines. At high pH (greater than its pKa of ~6.3) the amines are deprotonated and chitosan becomes insoluble, allowing for film formation. Following deposition, the films will remain attached to the electrode. However, the ability exists to delaminate them if desired. The films will remain stable as long as the pH of the solution does not drop below the pKa. Acidic solutions protonate the amines and the subsequent electrostatic repulsions swell the gel until it dissolves 18. That is, the assembly/disassembly process is reversible on demand and allows for removal of deposited films and reuse of electrodes. Conveniently, the pH range at which the sol-gel transition takes place is close to that in which most biological components function optimally. This makes the process ideal for the retention of functionality during assembly6.
Alginate film formation is facilitated by the anodic electrolysis of water as well as the presence of calcium carbonate 7. The localized low pH at the anode solubilizes the calcium carbonate leading to the release calcium cations. These ions are chelated by alginate, forming a crosslinked network on the electrode surface. Alginate films are notably reversible by competition for calcium ions from other chelating compounds such as citrate or EDTA, which can be used to dissolve the films, allowing for the re-use of the underlying electrodes. Thus, alginate films are relatively fragile when subjected to physiological conditions because calcium ions are easily scavenged from the gel matrix, weakening its structure and promoting film delamination or redissolution. To overcome this limitation, we have included an incubation step for the film in 1 M CaCl2 to strengthen the gel. Additionally, we recommend that the film's incubation solution (cell media, etc.) be supplemented with CaCl2 at a concentration of 500 &mu;M-3 mM.
The second major procedure is the functionalization of the deposited film with relevant biological components. This can be achieved in two ways, the first being electrochemical conjugation, a strategy that allows for rapid, reagentless assembly of proteins with exceptional spatial control 10. However, functionalization in this manner is limited by the diffusion of Cl- ions through the film to the electrode as well as the diffusion of HOCl, the generated reactive intermediate, back out into solution. The ability of electrochemically active molecules to pass through the film allows for the transduction of chemical and biological signals into easy-to-read electrical signals 15. We have shown tyrosinase-mediated coupling as a second strategy for enzyme functionalization to chitosan, demonstrated by covalently attaching AI-2 Synthase. This strategy allows the functionalization process to be controlled and selective - dependent on a specific reagent, tyrosinase, which acts discriminately on proteins containing a tyrosine tag 9. 
We show the usefulness and biocompatibility of multi-address systems by replicating natural pathways on a chip. First we organized two cell populations (i.e. "senders" and "receivers") at distinct addresses, and showed that they interacted across adjacent electrodes to deliver AI-2 and generate a fluorescence response. This concept has also been demonstrated by Cheng et al. in a microfluidic chip 14. We also mimicked the interaction, but instead used an enzyme to synthesize AI-2 for delivery. In this way, a synthetic intracellular pathway, AI-2 synthesis, was replicated through biofabrication and functioned much as it would in solution.
In both cases, assembly of multiple addresses presents the challenge of avoiding non-specific binding between addresses because each deposition solution must be introduced to the entire electrode array, even though electrodeposition is only intended at one address. Gentle yet thorough washing can remove the majority of residual solution from non-biased electrodes; the use of flow in microfluidic channels may further minimize non-specificity. Particularly for the adjacent biofabrication of chitosan and alginate addresses, we recommend depositing the chitosan film first, following this with biofunctionalization steps, and after this, electrodepositing alginate. Although we have not done so here, we have found that blocking the chitosan film with inert proteins (such as milk, BSA, etc.) greatly diminishes non-specific binding of unwanted molecules to chitosan's aminated surface.
We have found utility in establishing patterned electrodes, often found in bioMEMS devices, as the "blueprints" for a complex arrangement of cells and biomolecules. The uses of electrodeposited chitosan in bioMEMS devices can go well beyond the examples mentioned here 19. Chitosan can be deposited on various microscale geometries - such as in microchannels and on non-planar surfaces 20,15. The films can also be modified with other polymers and a variety of proteins, DNA, nanoparticles, and redox-active molecules for novel properties 21,22,23. In bioMEMS devices, chitosan films have been used for drug delivery, redox and small molecule detection, biocatalysis, and cell studies 20,23,24,25. Similarly, alginate is widely used as a cell-entrapment matrix and has been explored for reversible fluidic containment of cell populations and in-film immunoanalysis 26,27,28. Composite films for tissue engineering applications have been fabricated using alginate electrodeposition, with components such as with hydroxyapatite for orthopedic implants 29. 
In our demonstrations of biofabrication, we have shown both the interactions between biological components and across the bio-electronic interface to be equally applicable; this brings into reach the prospect of integrating all varieties of interactions for sophisticated performance in on-chip signal transmission. Accordingly, biofabrication may facilitate the fabrication of devices with reduced "minimum feature sizes" as a direct follow-on to rapid developments in microfabrication, as often motivated by consumer electronics. That is, next next-generation devices might in fact include labile biological components that offer nature's exquisite assembly and recognition capabilities at even smaller length scales than man-made systems. We envision near-term applications in analytical instrumentation, environmental sensors, and even biocompatible implantable devices.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the component</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Power Supply</td>
 <td>Keithley</td>
 <td>SourceMeter 2400</td>
 </tr>
 <tr>
 <td>Three electrode potentiostat</td>
 <td>CH Instruments</td>
 <td>Potentiostat/Galvanostat 600D </td>
 </tr>
 <tr>
 <td>RE-5B Ag/AgCl Reference Electrode with Flexible Connector </td>
 <td>BASi</td>
 <td>MF-2052</td>
 </tr>
 <tr>
 <td>Gold coated silicon wafer, 500um Si, 12nM Cr, 120nM Au, SiO2 for insulation</td>
 <td> custom fabricated</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Indium Tin oxide coated glass slide, rectangular, 8-12 ohm resist</td>
 <td>Sigma-Aldrich</td>
 <td>578274 </td>
 </tr>
 <tr>
 <td>Platinum sheet/foil (0.002 in)</td>
 <td>Surepure Chemetals</td>
 <td>1897</td>
 </tr>
 <tr>
 <td>Slim Line 2" Alligator Clips </td>
 <td>RadioShack</td>
 <td>270-346 </td>
 </tr>
 <tr>
 <td>Multi-Stacking Banana Plug Patch Cord</td>
 <td>TSElectronic</td>
 <td>B-36-02 B-24-02</td>
 </tr>
 <tr>
 <td>SYLGARD 184 silicone elastomer kit</td>
 <td>Dow Corning</td>
 <td>NC9020938 From Fischer</td>
 </tr>
 <tr>
 <td>Fluorescecence stereomicroscope</td>
 <td>Olympus</td>
 <td>MVX10 MacroView</td>
 </tr>
 <tr>
 <td>cellSens Standard </td>
 <td>Olympus</td>
 <td>version 1.3</td>
 </tr>
 </tbody>
</table>
Table 1. Electrodeposition and fluorescence visualization equipment.
<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Chitosan, medium molecular weight</td>
 <td>Sigma-Aldrich</td>
 <td>448877</td>
 </tr>
 <tr>
 <td>Hydrochloric Acid, ARISTAR. ACS, NF, FCC Grade </td>
 <td>VWR</td>
 <td>BDH3030</td>
 </tr>
 <tr>
 <td>Sodium Hydroxide, Solution. 10.00N</td>
 <td>VWR</td>
 <td>VW3247</td>
 </tr>
 <tr>
 <td>Alginic acid, sodium salt</td>
 <td>Sigma-Aldrich</td>
 <td>180947</td>
 </tr>
 <tr>
 <td>Multifex-MM Precipitated Calcium Carbonate, 70nm particles</td>
 <td>Speciality Minerals Inc.</td>
 <td>100-3630-3</td>
 </tr>
 </tbody>
</table>
Table 2. Chitosan and alginate solution reagents.
<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Calcium chloride, dihydrate</td>
 <td>J.T. Baker</td>
 <td>0504 </td>
 </tr>
 <tr>
 <td>Sodium Chloride, Certified ACS crystalline</td>
 <td>Fischer Scientific</td>
 <td>S271</td>
 </tr>
 <tr>
 <td>Potassium Phosphate Monobasic, anhydrous</td>
 <td>Sigma-Aldrich</td>
 <td>P9791</td>
 </tr>
 <tr>
 <td>Potassium Phosphate Dibasic, anhydrous</td>
 <td>Sigma- Aldrich</td>
 <td>P3786</td>
 </tr>
 <tr>
 <td>Phosphate Buffered Saline</td>
 <td>Sigma- 
 Aldrich</td>
 <td>P4417</td>
 </tr>
 </tbody>
</table>
Table 3. Other solution components and buffer reagents.
<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company/Source</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Glucose oxidase from aspergillus niger</td>
 <td>Sigma-Aldrich</td>
 <td>G2133 </td>
 </tr>
 <tr>
 <td>Tyrosinase from mushroom</td>
 <td>Sigma-Aldrich</td>
 <td>T3824</td>
 </tr>
 <tr>
 <td>LB broth, Miller (granulated)</td>
 <td>Fischer Scientific</td>
 <td>BP9723-2</td>
 </tr>
 <tr>
 <td>"AI2-Synthase" (HGLPT)</td>
 <td>Lab stock 16 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>W3110 wildtype cells</td>
 <td>Lab stock 30 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>MDAI2 + pCT6-lsrR-ampr + pET-dsRed-kanr cells </td>
 <td>Lab stock 30 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>FluoroSpheres: 1&mu;m diameter, Ex/Em: 505/515</td>
 <td>Invitrogen</td>
 <td>F8765</td>
 </tr>
 <tr>
 <td>5-(and-6)-carboxyrhodamine 6G succinimidyl ester, Ex/Em: 525/560</td>
 <td>Invitrogen</td>
 <td>C-6157 </td>
 </tr>
 <tr>
 <td>DyLight antibody labeling kit, 405</td>
 <td>Thermo Scientific</td>
 <td>PI-53020</td>
 </tr>
 </tbody>
</table>
Table 4. Enzymes, cells, and other functionalization reagents.

bridging the bio-electronic interface with biofabrication

Advancements in lab-on-a-chip technology promise to revolutionize both research and medicine through lower costs, better sensitivity, portability, and higher throughput. The incorporation of biological components onto biological microelectromechanical systems (bioMEMS) has shown great potential for achieving these goals. Microfabricated electronic chips allow for micrometer-scale features as well as an electrical connection for sensing and actuation. Functional biological components give the system the capacity for specific detection of analytes, enzymatic functions, and whole-cell capabilities. Standard microfabrication processes and bio-analytical techniques have been successfully utilized for decades in the computer and biological industries, respectively. Their combination and interfacing in a lab-on-a-chip environment, however, brings forth new challenges. There is a call for techniques that can build an interface between the electrode and biological component that is mild and is easy to fabricate and pattern.
Biofabrication, described here, is one such approach that has shown great promise for its easy-to-assemble incorporation of biological components with versatility in the on-chip functions that are enabled. Biofabrication uses biological materials and biological mechanisms (self-assembly, enzymatic assembly) for bottom-up hierarchical assembly. While our labs have demonstrated these concepts in many formats 1,2,3, here we demonstrate the assembly process based on electrodeposition followed by multiple applications of signal-based interactions. The assembly process consists of the electrodeposition of biocompatible stimuli-responsive polymer films on electrodes and their subsequent functionalization with biological components such as DNA, enzymes, or live cells 4,5. Electrodeposition takes advantage of the pH gradient created at the surface of a biased electrode from the electrolysis of water 6,7,. Chitosan and alginate are stimuli-responsive biological polymers that can be triggered to self-assemble into hydrogel films in response to imposed electrical signals 8. The thickness of these hydrogels is determined by the extent to which the pH gradient extends from the electrode. This can be modified using varying current densities and deposition times 6,7. This protocol will describe how chitosan films are deposited and functionalized by covalently attaching biological components to the abundant primary amine groups present on the film through either enzymatic or electrochemical methods 9,10. Alginate films and their entrapment of live cells will also be addressed 11. Finally, the utility of biofabrication is demonstrated through examples of signal-based interaction, including chemical-to-electrical, cell-to-cell, and also enzyme-to-cell signal transmission. 
Both the electrodeposition and functionalization can be performed under near-physiological conditions without the need for reagents and thus spare labile biological components from harsh conditions. Additionally, both chitosan and alginate have long been used for biologically-relevant purposes 12,13. Overall, biofabrication, a rapid technique that can be simply performed on a benchtop, can be used for creating micron scale patterns of functional biological components on electrodes and can be used for a variety of lab-on-a-chip applications.

Watch this Scientific Journal Video about Bridging the Bio-Electronic Interface with Biofabrication at JoVE.com

Bridging the Bio-Electronic Interface with Biofabrication

This article describes a biofabrication approach: deposition of stimuli-responsive polysaccharides in the presence of biased electrodes to create biocompatible films which can be functionalized with cells or proteins. We demonstrate a bench-top strategy for the generation of the films as well as their basic uses for creating interactive biofunctionalized surfaces for lab-on-a-chip applications.

Advancements in lab-on-a-chip technology promise to revolutionize both research and medicine through lower costs, better sensitivity, portability, and ...

bridging-the-bio-electronic-interface-with-biofabrication

Research

JoVE Journal

Bioengineering

Adhesion Frequency Assay for In Situ Kinetics Analysis of Cross-Junctional Molecular Interactions at the Cell-Cell Interface

The micropipette adhesion assay was developed in 1998 to measure two-dimensional (2D) receptor-ligand binding kinetics1. The assay uses a human red blood cell (RBC) as adhesion sensor and presenting cell for one of the interacting molecules. It employs micromanipulation to bring the RBC into contact with another cell that expresses the other interacting molecule with precisely controlled area and time to enable bond formation. The adhesion event is detected as RBC elongation upon pulling the two cells apart. By controlling the density of the ligands immobilized on the RBC surface, the probability of adhesion is kept in mid-range between 0 and 1. The adhesion probability is estimated from the frequency of adhesion events in a sequence of repeated contact cycles between the two cells for a given contact time. Varying the contact time generates a binding curve. Fitting a probabilistic model for receptor-ligand reaction kinetics1 to the binding curve returns the 2D affinity and off-rate.
The assay has been validated using interactions of Fc&gamma; receptors with IgG Fc1-6, selectins with glycoconjugate ligands6-9, integrins with ligands10-13, homotypical cadherin binding14, T cell receptor and coreceptor with peptide-major histocompatibility complexes15-19.
The method has been used to quantify regulations of 2D kinetics by biophysical factors, such as the membrane microtopology5, membrane anchor2, molecular orientation and length6, carrier stiffness9, curvature20, and impingement force20, as well as biochemical factors, such as modulators of the cytoskeleton and membrane microenvironment where the interacting molecules reside and the surface organization of these molecules15,17,19.
The method has also been used to study the concurrent binding of dual receptor-ligand species3,4, and trimolecular interactions19 using a modified model21.
The major advantage of the method is that it allows study of receptors in their native membrane environment. The results could be very different from those obtained using purified receptors17. It also allows study of the receptor-ligand interactions in a sub-second timescale with temporal resolution well beyond the typical biochemical methods.
To illustrate the micropipette adhesion frequency method, we show kinetics measurement of intercellular adhesion molecule 1 (ICAM-1) functionalized on RBCs binding to integrin &alpha;L&beta;2 on neutrophils with dimeric E-selectin in the solution to activate &alpha;L&beta;2.

Adhesion Frequency Assay for In Situ Kinetics Analysis of Cross-Junctional Molecular Interactions at the Cell-Cell Interface

An adhesion frequency assay for measuring receptor-ligand interaction kinetics when both molecules are anchored on the surfaces of the interacting cells is described. This mechanically-based assay is exemplified using a micropipette-pressurized human red blood cell as adhesion sensor and integrin &alpha;L&beta;2 and intercellular adhesion molecule-1 as interacting receptors and ligands.

The micropipette adhesion assay was developed in 1998 to measure two-dimensional (2D) receptor-ligand binding kinetics1. The assay uses a human red blood ...

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 ...

Development of Amelogenin-chitosan Hydrogel for In Vitro Enamel Regrowth with a Dense Interface

Biomimetic enamel reconstruction is a significant topic in material science and dentistry as a novel approach for the treatment of dental caries or erosion. Amelogenin has been proven to be a critical protein for controlling the organized growth of apatite crystals. In this paper, we present a detailed protocol for superficial enamel reconstruction by using a novel amelogenin-chitosan hydrogel. Compared to other conventional treatments, such as topical fluoride and mouthwash, this method not only has the potential to prevent the development of dental caries but also promotes significant and durable enamel restoration. The organized enamel-like microstructure regulated by amelogenin assemblies can significantly improve the mechanical properties of etched enamel, while the dense enamel-restoration interface formed by an in situ regrowth of apatite crystals can improve the effectiveness and durability of restorations. Furthermore, chitosan hydrogel is easy to use and can suppress bacterial infection, which is the major risk factor for the occurrence of dental caries. Therefore, this biocompatible and biodegradable amelogenin-chitosan hydrogel shows promise as a biomaterial for the prevention, restoration, and treatment of defective enamel.

Development of Amelogenin-chitosan Hydrogel for In Vitro Enamel Regrowth with a Dense Interface

In this article, we describe a protocol for fabricating an amelogenin-chitosan hydrogel for superficial enamel reconstruction. Organized in situ growth of apatite crystals in the hydrogel formed a dense enamel-restoration interface, which will improve the effectiveness and durability of restorations.

Biomimetic enamel reconstruction is a significant topic in material science and dentistry as a novel approach for the treatment of dental caries or ...

Methods for the Self-integration of Megamolecular Biopolymers on the Drying Air-LC Interface

Living organisms that use water are always prone to drying in the environment. Their activities are driven by biopolymer-based micro- and macro-structures, as seen in the cases of moving water in vascular bundles and moisturizing water in skin layers. In this study, we developed a method for assessing the effect of aqueous liquid crystalline (LC) solutions composed of biopolymers on drying. As LC biopolymers have megamolecular weight, we chose to study polysaccharides, cytoskeletal proteins, and DNA. The observation of biopolymer solutions during drying under polarized light reveals milliscale self-integration starting from the unstable air-LC interface. The dynamics of the aqueous LC biopolymer solutions can be monitored by evaporating water from a one-side-open cell. By analyzing the images taken using cross-polarized light, it is possible to recognize the spatio-temporal changes in the orientational order parameter. This method can be useful for the characterization of not only artificial materials in various fields, but also natural living tissues. We believe that it will provide an evaluation method for soft materials in the biomedical and environmental fields.

A method for the drying-induced self-integration of megamolecular biopolymers on the air-liquid crystalline interface is provided here. This methodology will be useful not only for understanding the macroscopic potentials of biopolymers, but also as an evaluation method for soft materials in biomedical and environmental fields.

Living organisms that use water are always prone to drying in the environment. Their activities are driven by biopolymer-based micro- and ...

Using Synthetic Biology to Engineer Living Cells That Interface with Programmable Materials

We have developed an abiotic-biotic interface that allows engineered cells to control the material properties of a functionalized surface. This system is made by creating two modules: a synthetically engineered strain of E. coli cells and a functionalized material interface. Within this paper, we detail a protocol for genetically engineering selected behaviors within a strain of E. coli using molecular cloning strategies. Once developed, this strain produces elevated levels of biotin when exposed to a chemical inducer. Additionally, we detail protocols for creating two different functionalized surfaces, each of which is able to respond to cell-synthesized biotin. Taken together, we present a methodology for creating a linked, abiotic-biotic system that allows engineered cells to control material composition and assembly on nonliving substrates.

This paper presents a series of protocols for developing engineered cells and functionalized surfaces that enable synthetically engineered E. coli to control and manipulate programmable material surfaces.

We have developed an abiotic-biotic interface that allows engineered cells to control the material properties of a functionalized surface. This system is ...

Fabrication of the Composite Regenerative Peripheral Nerve Interface (C-RPNI) in the Adult Rat

Recent advances in neuroprosthetics have enabled those living with extremity loss to reproduce many functions native to the absent extremity, and this is often accomplished through integration with the peripheral nervous system. Unfortunately, methods currently employed are often associated with significant tissue damage which prevents prolonged use. Additionally, these devices often lack any meaningful degree of sensory feedback as their complex construction dampens any vibrations or other sensations a user may have previously depended on when using more simple prosthetics. The composite regenerative peripheral nerve interface (C-RPNI) was developed as a stable, biologic construct with the ability to amplify efferent motor nerve signals while providing simultaneous afferent sensory feedback. The C-RPNI consists of a segment of free dermal and muscle graft secured around a target mixed sensorimotor nerve, with preferential motor nerve reinnervation of the muscle graft and sensory nerve reinnervation of the dermal graft. In rats, this construct has demonstrated the generation of compound muscle action potentials (CMAPs), amplifying the target nerve&#39;s signal from the micro- to milli-volt level, with signal to noise ratios averaging approximately 30-50. Stimulation of the dermal component of the construct generates compound sensory nerve action potentials (CSNAPs) at the proximal nerve. As such, this construct has promising future utility towards the realization of the ideal, intuitive prosthetic.

Fabrication of the Composite Regenerative Peripheral Nerve Interface C-RPNI in the Adult Rat

The following manuscript describes a novel method for developing a biologic, closed loop neural feedback system termed the composite regenerative peripheral nerve interface (C-RPNI). This construct has the ability to integrate with peripheral nerves to amplify efferent motor signals while simultaneously providing afferent sensory feedback.

Recent advances in neuroprosthetics have enabled those living with extremity loss to reproduce many functions native to the absent extremity, and this is ...

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 ...

Temperature-Controlled Assembly and Characterization of a Droplet Interface Bilayer

The droplet interface bilayer (DIB) method for assembling lipid bilayers (i.e., DIBs) between lipid-coated aqueous droplets in oil offers key benefits versus other methods: DIBs are stable and often long-lasting, bilayer area can be reversibly tuned, leaflet asymmetry is readily controlled via droplet compositions, and tissue-like networks of bilayers can be obtained by adjoining many droplets. Forming DIBs requires spontaneous assembly of lipids into high density lipid monolayers at the surfaces of the droplets. While this occurs readily at room temperature for common synthetic lipids, a sufficient monolayer or stable bilayer fails to form at similar conditions for lipids with melting points above room temperature, including some cellular lipid extracts. This behavior has likely limited the compositions&#8212;and perhaps the biological relevance&#8212;of DIBs in model membrane studies. To address this problem, an experimental protocol is presented to carefully heat the oil reservoir hosting DIB droplets and characterize the effects of temperature on the lipid membrane. Specifically, this protocol shows how to use a thermally conductive aluminum fixture and resistive heating elements controlled by a feedback loop to prescribe elevated temperatures, which improves monolayer assembly and bilayer formation for a wider set of lipid types. Structural characteristics of the membrane, as well as the thermotropic phase transitions of the lipids comprising the bilayer, are quantified by measuring the changes in electrical capacitance of the DIB. Together, this procedure can aid in evaluating biophysical phenomena in model membranes over various temperatures, including determining an effective melting temperature (TM) for multi-component lipid mixtures. This capability will thus allow for closer replication of natural phase transitions in model membranes and encourage the formation and use of model membranes from a wider swath of membrane constituents, including those that better capture the heterogeneity of their cellular counterparts.

This protocol details the use of a feedback temperature-controlled heating system to promote lipid monolayer assembly and droplet interface bilayer formation for lipids with elevated melting temperatures, and capacitance measurements to characterize temperature-driven changes in the membrane.

The droplet interface bilayer (DIB) method for assembling lipid bilayers (i.e., DIBs) between lipid-coated aqueous droplets in oil offers key benefits ...

The Fabrication and Operation of a Continuous Flow, Micro-Electroporation System with Permeabilization Detection

Current therapeutic innovations, such as CAR-T cell therapy, are heavily reliant on viral-mediated gene delivery. Although efficient, this technique is accompanied by high manufacturing costs, which has brought about an interest in using alternative methods for gene delivery. Electroporation is an electro-physical, non-viral approach for the intracellular delivery of genes and other exogenous materials. Upon the application of an electric field, the cell membrane temporarily allows molecular delivery into the cell. Typically, electroporation is performed on the macroscale to process large numbers of cells. However, this approach requires extensive empirical protocol development, which is costly when working with primary and difficult-to-transfect cell types. Lengthy protocol development, coupled with the requirement of large voltages to achieve sufficient electric-field strengths to permeabilize the cells, has led to the development of micro-scale electroporation devices. These micro-electroporation devices are manufactured using common microfabrication techniques and allow for greater experimental control with the potential to maintain high throughput capabilities. This work builds off a microfluidic-electroporation technology capable of detecting the level of cell membrane permeabilization at a single-cell level under continuous flow. However, this technology was limited to 4 cells processed per second, and thus a new approach for increasing the system throughput is proposed and presented here. This new technique, denoted as cell-population-based feedback control, considers the cell permeabilization response to a variety of electroporation pulsing conditions and determines the best-suited electroporation pulse conditions for the cell type under test. A higher-throughput mode is then used, where this 'optimal' pulse is applied to the cell suspension in transit. The steps for fabricating the device, setting up and running the microfluidic experiments, and analyzing the results are presented in detail. Finally, this micro-electroporation technology is demonstrated by delivering a DNA plasmid encoding for green fluorescent protein (GFP) into HEK293 cells.

This protocol describes the microfabrication techniques required to build a lab-on-a-chip, microfluidic electroporation device. The experimental setup performs controlled, single-cell-level transfections in a continuous flow and can be extended to higher throughputs with population-based control. An analysis is provided showcasing the ability to electrically monitor the degree of cell membrane permeabilization in real-time.

Current therapeutic innovations, such as CAR-T cell therapy, are heavily reliant on viral-mediated gene delivery. Although efficient, this technique is ...