Name: using synthetic biology to engineer living cells that interface with programmable materials
Uploaded: 2017-03-09T15:00:00
Description: Watch this Scientific Journal Video about Using Synthetic Biology to Engineer Living Cells That Interface with Programmable Materials at JoVE.com

The authors gratefully acknowledge support from award FA9550-13-1-0108 from the Air Force Office of Scientific Research of the USA. The authors additionally acknowledge support from award N00014-15-1-2502 from the Office of Naval Research of the USA, funding from the Institute for Critical Technology and Applied Science at Virginia Polytechnic Institute and State University, and from the National Science Foundation Graduate Research Fellowship Program, award number 1607310.

1. Media and Culture Preparation
<ol>
	<li>Prepare lysogeny broth (LB) media by mixing 25 g of LB powder stock with 1 L of deionized (DI) water and autoclaving the solution at 121 &#176;C for 20 min to sterilize.
	<ol>
		<li>To prepare LB plates, add 15 g agar (1.5%) to the LB media before sterilization</li>
		<li>Prepare stock solutions of 1,000x carbenicillin (Cb) in DI water (50 mg/mL).</li>
		<li>If preparing LB media that includes an antibiotic for selection of resistant transformants, wait until ...

<ol>
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We have presented a new strategy for interfacing engineered living cells with a functionalized material surface. This was accomplished by developing a cell line capable of synthesizing elevated levels of biotin when induced with IPTG. The elevated levels of biotin may then be used to modify the functionalized surface. The protocols detailed how to engineer the E. coli cell line and how to create two different functionalized surfaces.
Critical steps in this protocol occur throughout th...

Here, we report the procedures for developing a programmable substrate capable of responding to a chemical signal from an engineered cell line.1 We do this by creating a biotin-streptavidin interface that responds to biotin produced by synthetically engineered Escherichia coli (E. coli) cells. Previously, programmable surfaces have been engineered for a wide range of applications from toxin detection2 and point-of-care diagnosis3 to defense and security.4 While programmable surfaces can be useful as sensors and actuators, they can be made &#34;smarter&#34; by endowing them with the ability to adapt to different environmental challenges. In contrast, even simple microorganisms, such as E. coli, have inherent adaptability and are capable of responding to challenges with sophisticated and often unexpected solutions. This adaptability has enabled E. coli populations, controlled by their complex gene networks, to cost-effectively seek resources,5 create value-added products,6 and even power micro-scale robotics.7 By coupling the adaptive advantages of living cells with the use of programmable surfaces, we can create a smart substrate capable of responding to different environmental conditions.
Synthetic biology has given researchers new abilities to program the behavior of living organisms. By engineering cells to contain new gene regulatory networks, researchers can design cells that exhibit a range of programmed behaviors.8,9 Beyond basic research, these behaviors may be used for applications such as controlling material assembly and biologically producing value-added products.10 Herein, we detail how we used the tools of synthetic biology to engineer an E. coli strain that synthesizes biotin upon induction. This strain was developed by using restriction enzyme cloning methods to assemble a plasmid, pKE1-lacI-bioB. This plasmid, when transformed into E. coli strain K-12 MG1655, endows cells with the ability to express elevated levels of bioB, an essential enzyme for biotin synthesis. When transformed cells were induced with isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) and provided with a biotin precursor, desthiobiotin (DTB), elevated levels of biotin were produced.
Biotin&#39;s binding interaction with streptavidin is one of the strongest non-covalent bonds found in nature. As such, the biotin-streptavidin interaction is both well-characterized and highly employed in biotechnology.11 Within this manuscript, we present two strategies employing the biotin-streptavidin interaction to sense and detect cell-produced biotin with a functionalized surface. We refer to these contrasting surfaces as &#34;indirect&#34; and &#34;direct&#34; control schemes. In the indirect control scheme, cell-produced biotin competes with biotin that has been conjugated and immobilized on a polystyrene surface for streptavidin binding sites. In addition, the streptavidin is conjugated with horseradish peroxidase (HRP). HRP modifies 3, 3&#39;, 5, 5&#39;-tetramethylbenzidine (TMB), to produce an optical signal,12 which may be monitored by quantifying the spectral absorbance (i.e., optical density) at 450 nm (OD450). Thus, the indirect control scheme allows researchers to measure cell-produced biotin by monitoring the attentuation of the OD450 signal.
The direct control scheme exploits the streptavidin-biotin event by immobilizing streptavidin directly to a material surface and allowing cell-produced biotin and biotinylated HRP to compete for streptavidin binding sites. Again, the relative levels of cell-produced biotin are monitored by measuring an OD450 signal.
Taken together, the engineered cells and functionalized surfaces allow us to control the properties of a programmable surface by inducing networks in living cells. In other words, we have created a system that takes advantage of the adaptability of living organisms and the reliability and specification of an engineered material interface by linking these systems together.

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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.

Representative results are presented in the accompanying five figures. First, we present the cloning process graphically (Figure 1) so that the reader can visually follow the critical steps for creating the synthetically engineered E. coli strain. In order to characterize the population dynamics of the cells, we provide a growth curve (Figure 2) generated by measuring the optical density at 600 nm (OD600) of the population. Then, we sh...

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

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 ...

The overall goal of these procedures is to use synthetically engineered E.coli to control and manipulate programmable material surfaces by combining genetic modification and surface functionalization strategies. This method can help answer key questions in fields such as molecular medicine and environmental monitoring. By using genetically programmed cells to interpret a local environment and modify a functionalized material accordingly, we are providing a modular tool for a wide variety of applications.The main advantage of this technique is that living cells are able to act as dynamic sensors capable of reading, processing and recording the conditions around them via functionalized interfaces. After preparing solutions and collecting the biotin-enriched supernatant from biotin-producing E.coli according to the text protocol add 1.4 microliters of SPDP solution to 20 microliters of streptavidin or SA solution. Wrap the tube in aluminum foil and incubate it at room temperature for one and a half hours to allow the SPDP cross linker to bond to SA via an amino group, forming a pyridyldithio-activated SA.Following the incubation, add 2.4 microliters of DTT solution to the tube and incubate the sample at room temperature for one hour to allow a pyridine-2-thione cleavage resulting in a sulfhydryl-activated SA.Next, add 7.5 microliters of SCC solution to 72 microliters of HRP solution, wrap it in aluminum foil, and incubate it at room temperature for one and a half hours.This results in a maleimide-activated HRP bound by an amino group. Mix 17 microliters of LC-LC biotin solution with 200 microliters of BSA solution, wrap the tube in foil and incubate the sample at room temperature for one and a half hour. This step allows biotin to conjugate to BSA via an amide bond.Transfer the SA-SPDP solution, the HRP-SMCC and the BSA-conjugated biotin solution to separate centrifugal concentrators in spin tubes. Centrifuge the SA-SPDP at 10, 000 times g until the tube volume reaches 100 microliters or for 16 minutes, then fill the spin tube to 500 microliters using PBS-EDTA before repeating the spin and the addition of PBS-EDTA five more times. After spinning the solution down to 100 microliters, store the stock at four degrees Celsius.For the HRP SMCC, centrifuge the tubes at 10, 000 times g until the volume reaches 25 microliters or for 16 minutes. Fill the spin tube to 4500 microliters using PBS before repeating the spin and the addition of PBS five more times. After spinning the solution down to 100 microliters, store the stock at four degrees Celsius.For the BSA biotin, centrifuge the sample at 10, 000 times g until the volume reaches 100 microliters or for 12 minutes, then fill the spin tube to 500 microliters using PBS. After repeating the spin and the addition of PBS four more times, centrifuge the tube until the volume reaches 100 microliters or for 12 minutes, then add 100 microliters of PBS to the tube and store the stock at four degrees Celsius. Next, combine 25 microliters of the 10 milligram per milliliter HRP with 25 microliters of 10 milligram per milliliter SA solution and store the tube at four degrees Celsius overnight, then prepare two aliquots of BSA biotin in PBS by adding 10 microliters of BSA biotin stock to 490 microliters of PBS.Pipette 100 microliters of the solution into each well of a 96 well plate. Wrap the plate in foil and incubate it at 37 degrees Celsius for one hour. Next, add 200 microliters of 05%Tween 80 and BPS to the wells.Incubate the plate at room temperature for two minutes, then aspirate and decant the liquid. After repeating the wash three times, add 200 microliters of the 0.5%casing solution to each of the wells, then incubate the plate at 37 degrees Celsius for an hour. Following another round of washes as before, mix 12 milliliters of 05%casing solution with 7.5 microliters of 05%Tween 80, then use this solution to dilute the SA-HRP stock solution one to 10, 000.Pipette 80 microliters of the diluted SA-HRP into each well of a 96 well plate, then add 20 microliters of the prepared biotin sample to each well of the plate. Incubate the plate at 37 degrees Celsius for one hour. This step allows for competitive binding between free biotin and immobilized BSA biotin for SA-HRP binding sites.Then after washing the plate three times as before, prepare a 20 milliliter solution of 50 millimolar sodium acetate, 1%TMB and 3%hydrogen peroxide at a 1, 000 to 10 to one ratio. Add 200 microliters of the solution to each well of the plate, wrap it in foil and incubate it at room temperature for 15 minutes. Pipette 50 microliters of two molar sulfuric acid into each well to stop the reaction, then use a plate reader to measure the OD 450.Following the preparation of solutions according to the text protocol, add 7.5 microliters of LC-LC biotin to 72 microliters of HRP. Wrap the solution in foil and incubate the sample at room temperature for one and a half hours with agitation. This step causes biotin to conjugate to HRP via an amide bond.Transfer the solution to a centrifugal concentrator within a spin tube and centrifuge the solution at 10, 000 times g until the volume reaches 100 microliters or for 12 minutes, then use PBS to fill the spin tube to 500 microliters and repeat the centrifugation and buffer addition four times. Next, add 100 microliters of 0.17 micrograms per milliliter SA in PBS to each well of a 96 well plate. Wrap the plate in foil and incubate it at 37 degrees Celsius for an hour.To wash the wells of the plate, pipette 200 microliters of 05%Tween 80 into each well. Incubate the plate at room temperature for two minutes and decant the liquid. After repeating the wash three more times, add 200 microliters of the 0.5%casing solution to the wells.Incubate the plate at 37 degrees Celsius for one hour before washing the wells three times as before. Using the washed and concentrated biotin HRP solution previously prepared, dilute the biotin HRP stock solution one to 10, 000, then add 80 microliters of the solution to the wells of a 96 well plate. Add 20 microliters of prepared biotin sample to each well of the plate and incubate the plate at 37 degrees Celsius for one hour to allow competitive binding between free biotin and biotin HRP for the immobilized SA binding sites.After washing the wells three times as before, prepare a 20 milliliter solution of 50 millimolar sodium acetate, 1%TMV, and 3%hydrogen peroxide at a 1, 000 to 10 to one ratio. Add 200 microliters of the solution to each well of the plate, cover it with foil, and incubate it at room temperature for 15 minutes, then add 50 microliters of two molar sulfuric acid to each well to stop the reaction. Finally, use a plate reader to measure the OD 450.The engineered inducible E.coli MG1655 wild type cells were grown and monitored in minimal medium as well as minimal medium supplemented with DTB and/or IPTG. These plots shows the OD 600 reading measured every five minutes for 24 hours. In this experiment, the fluorescent protein mCherry was used in place of the bioB gene so the induction profile could be measured when engineered cells were induced with IPTG.These results demonstrate the efficacy of the inducible gene network. Plotted here are the optical signals at OD 450 in response to varying concentrations of biotin for both the indirect and direct functionalized surface schemes. In this experiment, products of induced engineered cells were used to chemically modify the functionalized surface.The different biotin concentrations are presented as measured by the indirect control scheme functionalized surface for wild type cells, uninduced cells containing the pKE1-lacI-bioB plasmid and induced cells containing the pKE1-lacI-bioB plasmid. Once mastered, cells can be genetically engineered in two days, while the direct and indirect surface functionalization techniques can be each done in five hours if they are performed properly. While attempting this procedure, it's important to remember to avoid contamination of bacterial cell culture and cover the surfaces during functionalization to avoid photo bleaching.After watching this video, you should have a good understanding of how to functionalize surfaces for both the direct and indirect control schemes. Don't forget that working with ethidium bromide can be extremely hazardous, and precautions, such as wearing gloves, should always be taken while performing this procedure.

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Research

JoVE Journal

Bioengineering

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 ...

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.

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 ...

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, ...

Ratiometric Biosensors that Measure Mitochondrial Redox State and ATP in Living Yeast Cells

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell death. These processes affect and are affected by the redox status of and ATP production by mitochondria. Here, we describe the use of two ratiometric, genetically encoded biosensors that can detect mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells. Mitochondrial redox state is measured using redox-sensitive Green Fluorescent Protein (roGFP) that is targeted to the mitochondrial matrix. Mito-roGFP contains cysteines at positions 147 and 204 of GFP, which undergo reversible and environment-dependent oxidation and reduction, which in turn alter the excitation spectrum of the protein. MitGO-ATeam is a F&ouml;rster resonance energy transfer (FRET) probe in which the &epsilon; subunit of the FoF1-ATP synthase is sandwiched between FRET donor and acceptor fluorescent proteins. Binding of ATP to the &epsilon; subunit results in conformation changes in the protein that bring the FRET donor and acceptor in close proximity and allow for fluorescence resonance energy transfer from the donor to acceptor.

We describe the use of two ratiometric, genetically encoded biosensors, which are based on GFP, to monitor mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells.

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell ...

Characterization Of Multi-layered Fish Scales (Atractosteus spatula) Using Nanoindentation, X-ray CT, FTIR, and SEM

The hierarchical architecture of protective biological materials such as mineralized fish scales, gastropod shells, ram&#8217;s horn, antlers, and turtle shells provides unique design principles with potentials for guiding the design of protective materials and systems in the future. Understanding the structure-property relationships for these material systems at the microscale and nanoscale where failure initiates is essential. Currently, experimental techniques such as nanoindentation, X-ray CT, and SEM provide researchers with a way to correlate the mechanical behavior with hierarchical microstructures of these material systems1-6. However, a well-defined standard procedure for specimen preparation of mineralized biomaterials is not currently available. In this study, the methods for probing spatially correlated chemical, structural, and mechanical properties of the multilayered scale of A. spatula using nanoindentation, FTIR, SEM, with energy-dispersive X-ray (EDX) microanalysis, and X-ray CT are presented.

Characterization Of Multi-layered Fish Scales Atractosteus spatula Using Nanoindentation, X-ray CT, FTIR, and SEM

This paper presents the methods used for probing spatially correlated chemical, structural, and mechanical properties of the multilayered scale of Atractosteus spatula (A. spatula) using nanoindentation, Fourier transform infrared (FTIR) spectroscopy, scanning electron microscopy (SEM), and X-ray computed tomography (X-ray CT). The experimental results have been used to investigate the design principles of protective biological materials.

The hierarchical architecture of protective biological materials such as mineralized fish scales, gastropod shells, ram&#8217;s horn, antlers, and turtle ...

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 ...

Sealable Femtoliter Chamber Arrays for Cell-free Biology

Cell-free systems provide a flexible platform for probing specific networks of biological reactions isolated from the complex resource sharing (e.g., global gene expression, cell division) encountered within living cells. However, such systems, used in conventional macro-scale bulk reactors, often fail to exhibit the dynamic behaviors and efficiencies characteristic of their living micro-scale counterparts. Understanding the impact of internal cell structure and scale on reaction dynamics is crucial to understanding complex gene networks. Here we report a microfabricated device that confines cell-free reactions in cellular scale volumes while allowing flexible characterization of the enclosed molecular system. This multilayered poly(dimethylsiloxane) (PDMS) device contains femtoliter-scale reaction chambers on an elastomeric membrane which can be actuated (open and closed). When actuated, the chambers confine Cell-Free Protein Synthesis (CFPS) reactions expressing a fluorescent protein, allowing for the visualization of the reaction kinetics over time using time-lapse fluorescent microscopy. Here we demonstrate how this device may be used to measure the noise structure of CFPS reactions in a manner that is directly analogous to those used to characterize cellular systems, thereby enabling the use of noise biology techniques used in cellular systems to characterize CFPS gene circuits and their interactions with the cell-free environment.

A microfabricated device with sealable femtoliter-volume reaction chambers is described. This report includes a protocol for sealing cell-free protein synthesis reactants inside these chambers for the purpose of understanding the role of crowding and confinement in gene expression.

Cell-free systems provide a flexible platform for probing specific networks of biological reactions isolated from the complex resource sharing (e.g., ...

Creating a Structurally Realistic Finite Element Geometric Model of a Cardiomyocyte to Study the Role of Cellular Architecture in Cardiomyocyte Systems Biology

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal microscopy, the scientific community has unprecedented access to large datasets at sub-micrometer resolution that characterize the architectural remodeling that accompanies changes in cardiomyocyte function in health and disease. However, these datasets have been under-utilized for&#160;investigating the role of cellular architecture remodeling in cardiomyocyte function. The purpose of this protocol is to outline how to create an accurate finite element model of a cardiomyocyte using high resolution electron microscopy and confocal microscopy images. A detailed and accurate model of cellular architecture has significant potential to provide new insights into cardiomyocyte biology, more than experiments alone can garner. The power of this method lies in its ability to computationally fuse information from two disparate imaging modalities of cardiomyocyte ultrastructure to develop one unified and detailed model of the cardiomyocyte. This protocol outlines steps to integrate electron tomography and confocal microscopy images of adult male Wistar (name for a specific breed of albino rat) rat cardiomyocytes to develop a half-sarcomere finite element model of the cardiomyocyte. The procedure generates a 3D finite element model that contains an accurate, high-resolution depiction (on the order of ~35 nm) of the distribution of mitochondria, myofibrils and ryanodine receptor clusters that release the necessary calcium for cardiomyocyte contraction from the sarcoplasmic reticular network (SR) into the myofibril and cytosolic compartment. The model generated here as an illustration does not incorporate details of the transverse-tubule architecture or the sarcoplasmic reticular network and is therefore a minimal model of the cardiomyocyte. Nevertheless, the model can already be applied in simulation-based investigations into the role of cell structure in calcium signaling and mitochondrial bioenergetics, which is illustrated and discussed using two case studies that are presented following the detailed protocol.

This protocol outlines a novel method to create a spatially detailed finite element model of the intracellular architecture of cardiomyocytes from electron microscopy and confocal microscopy images. The power of this spatially detailed model is demonstrated using case studies in calcium signaling and bioenergetics.

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal ...

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 ...

Preparing Protein Producing Synthetic Cells using Cell Free Bacterial Extracts, Liposomes and Emulsion Transfer

The bottom-up assembly approach for construction of synthetic cells is an effective tool for isolating and investigating cellular processes in a cell mimicking environment. Furthermore, the development of cell-free expression systems has demonstrated the ability to reconstitute the protein production, transcription and translation processes (DNA&#8594;RNA&#8594;protein) in a controlled manner, harnessing synthetic biology. Here we describe a protocol for preparing a cell-free expression system, including the production of a potent bacterial lysate and encapsulating this lysate inside cholesterol-rich lipid-based giant unilamellar vesicles (GUVs) (i.e., stable liposomes), to form synthetic cells. The protocol describes the methods for preparing the components of the synthetic cells including the production of active bacterial lysates, followed by a detailed step-by-step preparation of the synthetic cells based on a water-in-oil emulsion transfer method. These facilitate the production of millions of synthetic cells in a simple and affordable manner with a high versatility for producing different types of proteins. The obtained synthetic cells can be used to investigate protein/RNA production and activity in an isolated environment, in directed evolution, and also as a controlled drug delivery platform for on-demand production of therapeutic proteins inside the body.

This protocol describes the method, materials, equipment and steps for bottom-up preparation of RNA and protein producing synthetic cells. The inner aqueous compartment of the synthetic cells contained the S30 bacterial lysate encapsulated within a lipid bilayer (i.e., stable liposomes), using a water-in-oil emulsion transfer method.

The bottom-up assembly approach for construction of synthetic cells is an effective tool for isolating and investigating cellular processes in a cell ...