The authors would like to extend their gratitude for financial support from the Pasteur Foundation of New York (to J.D., C.S., C.S.), the EU-FP7 Program "Automation" (to S.L.S.), the Institut Carnot Program 11 (to J.D., A.H., A.R., R.T., S.L.S.) and Project IMNOS (to R.T., S.L.S.), the Conny-Maeve Charitable Foundation (S.L.S.), the European Masters in Molecular Imaging (to I.T.), the Region Ile de France programs MODEXA (S.L.S.), SESAME (S.L.S.) and DimMalInf (S.L.S., R.T.), the ANR Program Grandes Investissement de l'avenir Infrastructures Nationales en Biologie-Sant&#233;: France LifebioImaging (FLI) France Life Imaging (R.T., S.L.S.), France Bioimaging (J.D., S.L.S) and the Institut Pasteur, Paris. Further, the authors would like to thank, Jos&#233; Bengoechea and Herbert Schweizer for reagents. Further, The authors would like to thank Cindy Fevre who generated the antibodies.

1. Reagents
<ol>
	<li>Purchase or develop luminescent bacteria and appropriate culture media such as modified Escherichia coli expressing the luxABCDE21, Vibrio fischeri, Photobacterium sp. Klebsiella pneumoniae22.</li>
	<li>Prepare physiological saline (0.9% NaCl) and culture media solutions according to standard recipes. E. coli and K. pneumoniae were grown in Luria Bertani (LB) at 37 &#176;C, and V. fischeri and Photobacterium sp in LB supplemented with 0.5 M NaCl at 22 &#176;C.</li>
	<li>Purchase or acquire fluorescent probes such as Q-tracker 705 (40 nm diameter, referred to as QD705), Q-tracker 800 (40 nm diameter, referred to as QD800), fluorescent (40 nm diameter) and non-fluorescent polystyrene microspheres (48 nm diameter), and conventional fluorescent dyes.</li>
	<li>Prepare the necessary reagents for bacterial labeling.</li>
	<li>Ensure access to a whole animal bioluminescence imager, such as an IVIS Spectrum, that is capable of detecting signal under a wide range of emission wavelengths and exposure times. A plate reader capable of bioluminescence measurements will suffice for most of the experiments described here.</li>
</ol>
2. Basic Recapitulation of FUEL
<ol>
	<li>Starting from individual colonies on standard culture plates, start overnight cultures of luminescent bacteria. Here, V. fischeri were used.</li>
	<li>The day of experimentation, initiate fresh subcultures and allow them to progress until an OD600 of 1-1.5 is achieved. In order to produce comparable results it is best to use bacteria in similar growth states in which they produce intense signals. This can be assured by keeping the OD600 constant.</li>
	<li>Combine aliquots of 100 &#956;l from each with either 5&#160;&#956;l of QD705 or physiological saline (PS), and then add to 895 &#956;l of PS into standard spectroscopic cuvettes. Place the filled cuvettes into the IVIS Spectrum and take the measurements under the appropriate filter sets. Here, the 710-730 nm emission filter was used.</li>
</ol>
3. FUEL Over Varying Distances
<ol>
	<li>Fill two reduced volume (1 ml) plastic photometric cuvettes with either 50 &#956;l of QD705 or 97.2 &#956;l of non-fluorescent 48 nm polystyrene microspheres in a total volume of 1 ml PS. This ensures similar solid surface area per total volume between the two entities.</li>
	<li>Prepare a light source cuvette by encasing a third cuvette with standard black tape or some other opaque material capable of blocking light. Carefully prepare two identical optical windows on opposite sides of the cuvette.</li>
	<li>Place the previously filled cuvettes directly onto either side of the light source cuvette. Add a 1 ml aliquot of V. fischeri or other culture (i.e. luminescent E. coli or Photobacterium sp) from a fresh subculture into the light source cuvette and cover it with an opaque material, such as black paper to reduce any light contamination.</li>
	<li>Visualize the three cuvettes under the appropriate emission filters such as the Total Light and 710-730 nm emission filters, with exposure times of 10, 30, and 30 sec, respectively.</li>
	<li>Reposition both external cuvettes at equivalent further distances from the central cuvette, and then visualize again. Repeat until a final face-to-face (central to reduced volume cuvette) distance of 3cm is achieved. At each step, acquire a fluorescence image (450-480 nm excitation, 710-730 nm emission) to validate the QD705 location.</li>
</ol>
4. Investigating Potential FUEL Pairs
<ol>
	<li>Fill two reduced volume cuvettes with 1 ml solutions containing either a fluorophore or a fluorescent nanoparticle of interest in one, and PS or PS with non-fluorescent nanoparticles as the negative control in the other. In the work demonstrated here, a variety of potential commercially available FUEL fluorophores and fluorescent nanoparticles were investigated as outlined in Table 1.</li>
	<li>Add a 1 ml aliquot of fresh V. fischeri or other luminescent bacteria to a blacked-out third cuvette containing two physically equal optical windows situated on opposite sides. Cover the cuvette with an opaque substance such as a piece of black paper.</li>
	<li>At a short but equal distance place the two reduced volume cuvettes on either side of the blacked-out cuvette and visualize under the appropriate filters. Here, a distance of 0.7 cm was used. Repeat with the other fluorophores.</li>
</ol>
5. Targeted FUEL
<ol>
	<li>Prepare biotinylated antibodies specific to the luminescent bacteria. For example, here primary antibodies specific to K. pneumoniae (&#945;-Kp) were biotinylated using a solution containing EZ-Link Sulfo-NHS-LC-Biotin.</li>
	<li>Remove excess reagent from the antibodies using desalting columns under centrifugation (1,500 x g) for 2 min.</li>
	<li>Use a standard Bradford assay to determine the protein concentration and determine the degree of antibody biotinylation by HABA assay.</li>
	<li>Obtain the &#945;-Kp-biot biotinylated antibody (Ab) batches by mixing 100 &#956;l (10 mM, dissolved in 1x PBS) with 40 &#956;l &#945;-Kp antibody, resulting in a final degree of substitution of 16 biotin residues per Ab and a final protein concentration of 10 mg ml-1.</li>
	<li>Using multiple overnight replicate cultures, target the washed bacteria with the aforementioned antibodies following the appropriate protocol.
	<ol>
		<li>Specifically, wash the K. pneumoniae in 1x PBS, and resuspend in 1x PBS to an OD of 4 (ca. 4 x 108 CFU ml-1 OD-1).</li>
		<li>For each replicate, incubate 100 &#956;l PBS, &#945;-Kp antibodies (10 &#956;l &#945;-Kp-biotB or 10 &#956;l PBS for the control) and 100 &#956;l cells for 90 min at 30 &#176;C. Wash the cells three times in 1x PBS, and resuspend in 196 &#956;l PBS.</li>
	</ol>
	</li>
	<li>Label the cells with the QD705 streptavidin conjugate and divide them into two equivalent volumes.</li>
	<li>Wash one solution three times and then resuspend it into the appropriate volume.</li>
	<li>Distribute 100 &#956;l of the washed and unwashed solutions into individual wells of a black 96-well plate. Measure the resulting luminescence under the Total Light, 490-510 nm, and 710-730 nm filters. Fluorophore concentration can be determined using 450-480 nm excitation and 710-730 nm emission.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioluminescence+imaging:+progress+and+applications.">Bioluminescence imaging: progress and applications.</a> Trends in Biotechnology. 29, 624-633 (2011).</li><li>Maywood, E. 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The authors declare competing financial interests. J.D., S.B., K.L.R. and S.L.S. contributed to European Patent EP10290158.4 and World Intellectual Property Organization Patent Application WO/2011/117847.

The fundamental demonstration of FUEL can be achieved simply by mixing luminescent bacteria with fluorescent nanoparticles or QDs. The two entities will be physically separated and remain beyond any efficient RET distance. More difficult is the FUEL signal optimization both in vitro and in vivo. Under in vitro conditions, both with and without an optical absorber present, usually the addition of excess fluorophore will be sufficient to maximize the FUEL response. However, at high concentrations phenomena such as static or collisional quenching can lead to a loss of fluorescent signal. Performing a dilution series by independently varying the concentration of the luminescent source and the fluorophore will help to optimize the desired concentrations. The establishment and optimization of FUEL under in vivo concentrations is much more difficult and needs to be addressed on a case-by-case basis. It can be difficult to create a condition where the fluorescent entity can be accessed optically by the luminescent source. As such, beginning with direct co-injections of the two moieties can provide information regarding the success of FUEL under optimal conditions.
Standard protocols exist for labeling bacteria and eukaryotic cells with fluorescent entities such as the Alexa series and QDs. Often this requires surface functionalization or activation with antibodies, which can lead to unwanted effects like reduced cell viability or altered metabolic activity. To overcome this, it is important to determine the optimal amount of antibody or activation agent needed that minimizes cellular perturbations while maximizing the fluorescent labeling. The use of QDs is advantageous because of their characteristically broad excitation spectra, narrow and tunable emission spectra, and the possibility of a large Stokes shift. However, QDs can be cytotoxic and may not be desirable in some cases.
FUEL is a phenomenon that is present in many BRET experiments13 and is applicable to a variety of luminescent and fluorescent sources. Until now, the photons resulting from FUEL were considered the product of non-specific interactions or an unfortunate background signal resulting from poorly designed BRET experiments. It is only with the type of experiments demonstrated here that we were able to identify the utility of this unwanted signal. In the shown examples, the luminescent bacteria act as a diffuse excitation source capable of eliciting a standard fluorescent response from a wide variety of fluorescent entities. Furthermore, due to the substantial working distance, it is safe to conclude that while FUEL can be constructed without the occurrence of BRET, in general BRET cannot be observed without a contribution from FUEL. Importantly, due to the lack of a targeting requirement, FUEL can be used to cover the spatial gap that exists between BRET and conventional whole-animal imaging techniques.

The genetic modification of organisms, such as viruses1,2, bacteria3, or small mammals4 to either induce or constitutively express bioluminescence, has been highly successful and widely demonstrated5-7. Bioluminescence, an in vivo chemiluminescent reaction involving naturally occurring reagents, has the advantage of producing light without the need for an external light source. As such, bioluminescent imaging does not suffer from the common drawbacks of auto- and non-specific signal found from fluorescence imaging8. Consequently, bioluminescence has a significant signal-to-noise ratio since any detected signal originates solely from the intended source. While many models have exploited the lux operon from Photorhabdus luminescens (emission maximum centered between 480 and 490 nm) for in vitro and in vivo applications9, its use in small mammals has been problematic due to the very nature of the imaging conditions; the pervading existence of optical absorbers, such as hemoglobin, and scattering agents, such as tissue and bone, strongly affect blue to yellow wavelengths3. The expression of an engineered firefly luciferase (emission maximum at 617nm) has been recently developed and incorporated, providing a tool that greatly overcomes optical absorption10, but is still subject to scattering effects.
In response, there have been multiple attempts to red-shift the emitted signal into the desired optical window of 650-900 nm, a region of minimized absorption and scatter, using bioluminescence resonance energy transfer (BRET)11-13. As a tool to enhance signal detection, BRET, which uses a bioluminescent source as the donor and an added fluorophore as the acceptor, has found limited success. As a seminal example of this phenomenon, &#34;self-illuminating quantum dots&#34; (SIQDs)14 consist of modified Renilla reniformis luciferases bound to the external polymer-lysine layer of commercially available quantum dots (QDs). Upon substrate addition, the resulting bioluminescent reaction induces fluorescence emission from the QDs, generating a significant production of red photons. However, these SIQDs have limited applicability to in vivo visualization of physiologically relevant events. This limited applicability is likely due to the difficulty of linking the dual probe to the organ, cell or gene of interest, since the SIQDs cannot be genetically encoded and therefore would require a secondary modification of the polymer shell. To improve their applicability, alternative SIQDs, where the luciferases are bound directly to the luminescent core, have recently been employed15. Building off of the SIQD concept, a more applicable BRET system was achieved by attaching Cypridina luciferase to an indocyanine dye16, which was capable of specifically targeting tumors in mice while producing a substantial red shift from 460 nm to 675 nm. To undergo non-radiative energy transfer, BRET follows the same primary constraints as its fluorescent counterpart: there must be a strong spectral overlap between the donor emission and the acceptor excitation spectra and the working distance between the two moieties must be on the order of the F&#246;rster radius (5-14 nm depending on the donor-acceptor pair, with an effective maximum distance of twice the F&#246;rster radius17). This distance dependence greatly limits the types of events that can be observed using BRET as a means to enhance detection.
Recently a new approach was identified and demonstrated under both in vitro and in vivo conditions. Building off the foundation of BRET, Fluorescence by Unbound Excitation from Luminescence (FUEL)18,19 also requires a strong spectral overlap between the luminescent and fluorescent components. However, unlike BRET, FUEL is a completely radiative process whereby the emitted photon from the luminescent source is absorbed by an optically accessible fluorophore, which subsequently emits a red-shifted photon according to the fluorophore quantum yield. Akin to BRET, this approach can also be used to overcome the constraints of imaging in the presence of optical absorbers. The resulting red shift provides an overall increase and specificity in the detected signal due to a decrease in attenuation and a reduction of optical scattering effects. FUEL has been reported to occur between bioluminescent Escherichia coli expressing the lux operon and QDs18,19. While experimentally similar to the SIQDs, a fundamental difference exists: in FUEL, it is not necessary for the luminescent source to be physically bound to the fluorophore, which allows for genetic encoding of the luminescent probe. Due to the successful detection of FUEL between luminescent bacteria and QDs, it is possible that this technique could be applied to both superficial (skin) and deep tissue (lung, liver) infections such as Staphylococcus aureus and Klebsiellia pneumoniae.
Since the report of its experimental significance, FUEL has evolved to include a robust mathematical model20 that can be used to predict acceptable luminescent and fluorescent pairs, and its applications have expanded to include use in identification of photophysical characteristics such as quantum yield. We describe below some of the basic techniques of FUEL. First, we show evidence for this phenomenon over both short (&#181;m) and long (cm) working distances, which fundamentally distinguishes FUEL from BRET. Second, we expand upon the possible FUEL pairs by examining a wide variety of fluorophores and fluorescent nanoparticles. Third, FUEL applications are investigated by comparing targeted and non-targeted FUEL pairs.

<table border="0" cellpadding="0" cellspacing="0" style="width:764px;" width="764">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Name of Material/ Equipment</td>
			<td style="width:175px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td colspan="2" height="21" style="height:21px;">Escherichia coli expressing the luxABCDE operon&#160;</td>
			<td style="width:119px;"></td>
			<td>kindly provided by Jos&#233; A. Bengoechea with permission from Herbert P. Schweizer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Klebsiella pneumoniae 52145&#160;</td>
			<td style="width:175px;"></td>
			<td style="width:119px;"></td>
			<td>52145 is a serotype K2 reference strain</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Luria Bertani (LB)&#160;</td>
			<td style="width:175px;"></td>
			<td style="width:119px;"></td>
			<td>standard growth media</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Q-Tracker 705&#160;</td>
			<td style="width:175px;">Life Sciences</td>
			<td>Q21061MP</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Q-Tracker 800</td>
			<td style="width:175px;">Life Sciences</td>
			<td>Q21071MP</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Alexa 555</td>
			<td style="width:175px;">Life Sciences</td>
			<td style="width:119px;">S21381</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Alexa 568</td>
			<td style="width:175px;">Life Sciences</td>
			<td style="width:119px;">S11226</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Alexa 633</td>
			<td style="width:175px;">Life Sciences</td>
			<td style="width:119px;">S21375</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Alexa 700</td>
			<td style="width:175px;">Life Sciences</td>
			<td style="width:119px;">S21383</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Non-fluorescent microspheres</td>
			<td style="width:175px;">Polysciences, Inc</td>
			<td align="right" style="width:119px;">15913</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Pink microspheres</td>
			<td style="width:175px;">Life Sciences</td>
			<td>F8887</td>
			<td>40nm diameter</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">yellow microspheres</td>
			<td style="width:175px;">Life Sciences</td>
			<td style="width:119px;">F8888</td>
			<td>40nm diameter</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Ivis Spectrum</td>
			<td style="width:175px;">PerkinElmer</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">EZ-Link Sulfo-NHS-LC-Biotin</td>
			<td style="width:175px;">Thermo Scientific Pierce</td>
			<td align="right">21425</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Zeba Spin Desalting Columns 7K MWCO</td>
			<td style="width:175px;">Thermo Scientific Pierce</td>
			<td align="right">21425</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HABA assay kit&#160;</td>
			<td style="width:175px;">Thermo Scientific Pierce</td>
			<td align="right">28005</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:304px;">Bradford assay</td>
			<td style="width:175px;">Bio-Rad</td>
			<td>500-0201</td>
			<td></td>
		</tr>
	</tbody>
</table>

a step beyond bret: fluorescence by unbound excitation from luminescence (fuel)

Fluorescence by Unbound Excitation from Luminescence (FUEL) is a radiative excitation-emission process that produces increased signal and contrast enhancement in vitro and in vivo. FUEL shares many of the same underlying principles as Bioluminescence Resonance Energy Transfer (BRET), yet greatly differs in the acceptable working distances between the luminescent source and the fluorescent entity. While BRET is effectively limited to a maximum of 2 times the F&ouml;rster radius, commonly less than 14 nm, FUEL can occur at distances up to &micro;m or even cm in the absence of an optical absorber. Here we expand upon the foundation and applicability of FUEL by reviewing the relevant principles behind the phenomenon and demonstrate its compatibility with a wide variety of fluorophores and fluorescent nanoparticles. Further, the utility of antibody-targeted FUEL is explored. The examples shown here provide evidence that FUEL can be utilized for applications where BRET is not possible, filling the spatial void that exists between BRET and traditional whole animal imaging.

Watch this Scientific Journal Video about A Step Beyond BRET: Fluorescence by Unbound Excitation from Luminescence FUEL at JoVE.com

A Step Beyond BRET: Fluorescence by Unbound Excitation from Luminescence FUEL

Expanding the foundation and applicability of Fluorescence by Unbound Excitation from Luminescence (FUEL) by surveying the relevant principles and demonstrating its compatibility with a multitude of fluorophores and antibody-targeted conditions.

A Step Beyond BRET: Fluorescence by Unbound Excitation from Luminescence (FUEL)

Fluorescence by Unbound Excitation from Luminescence (FUEL) is a radiative excitation-emission process that produces increased signal and contrast ...

a-step-beyond-bret-fluorescence-unbound-excitation-from-luminescence

Research

JoVE Journal

Bioengineering

11.5K Views.  Institut Pasteur.  توسيع المؤسسة وتطبيق الإسفار عن طريق الإثارة غير منضم من التلألؤ (FUEL) من خلال المسح إلى المبادئ ذات الصلة ومما يدل على توافقه مع العديد من fluorophores وظروف استهداف الأجسام المضادة. 

Video: A Step Beyond BRET: Fluorescence by Unbound Excitation from Luminescence FUEL

Thermodynamics of Membrane Protein Folding Measured by Fluorescence Spectroscopy

Membrane protein folding is an emerging topic with both fundamental and health-related significance. The abundance of membrane proteins in cells underlies the need for comprehensive study of the folding of this ubiquitous family of proteins. Additionally, advances in our ability to characterize diseases associated with misfolded proteins have motivated significant experimental and theoretical efforts in the field of protein folding. Rapid progress in this important field is unfortunately hindered by the inherent challenges associated with membrane proteins and the complexity of the folding mechanism. Here, we outline an experimental procedure for measuring the thermodynamic property of the Gibbs free energy of unfolding in the absence of denaturant, &Delta;G&deg;H2O, for a representative integral membrane protein from E. coli. This protocol focuses on the application of fluorescence spectroscopy to determine equilibrium populations of folded and unfolded states as a function of denaturant concentration. Experimental considerations for the preparation of synthetic lipid vesicles as well as key steps in the data analysis procedure are highlighted. This technique is versatile and may be pursued with different types of denaturant, including temperature and pH, as well as in various folding environments of lipids and micelles. The current protocol is one that can be generalized to any membrane or soluble protein that meets the set of criteria discussed below.

This video article details the experimental procedure for obtaining the Gibbs free energy of membrane protein folding by tryptophan fluorescence.

Membrane protein folding is an emerging topic with both fundamental and health-related significance.  The abundance of membrane proteins in cells ...

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 ...

Generation and Recovery of &beta;-cell Spheroids From Step-growth PEG-peptide Hydrogels

Hydrogels are hydrophilic crosslinked polymers that provide a three-dimensional microenvironment with tissue-like elasticity and high permeability for culturing therapeutically relevant cells or tissues. Hydrogels prepared from poly(ethylene glycol) (PEG) derivatives are increasingly used for a variety of tissue engineering applications, in part due to their tunable and cytocompatible properties. In this protocol, we utilized thiol-ene step-growth photopolymerizations to fabricate PEG-peptide hydrogels for encapsulating pancreatic MIN6 b-cells. The gels were formed by 4-arm PEG-norbornene (PEG4NB) macromer and a chymotrypsin-sensitive peptide crosslinker (CGGYC). The hydrophilic and non-fouling nature of PEG offers a cytocompatible microenvironment for cell survival and proliferation in 3D, while the use of chymotrypsin-sensitive peptide sequence (CGGY&darr;C, arrow indicates enzyme cleavage site, while terminal cysteine residues were added for thiol-ene crosslinking) permits rapid recovery of cell constructs forming within the hydrogel. The following protocol elaborates techniques for: (1) Encapsulation of MIN6 &beta;-cells in thiol-ene hydrogels; (2) Qualitative and quantitative cell viability assays to determine cell survival and proliferation; (3) Recovery of cell spheroids using chymotrypsin-mediated gel erosion; and (4) Structural and functional analysis of the recovered spheroids.

Generation and Recovery of &amp;#946;-cell Spheroids From Step-growth PEG-peptide Hydrogels

The following protocol provides techniques for encapsulating pancreatic &#x03B2;-cells in step-growth PEG-peptide hydrogels formed by thiol-ene photo-click reactions. This material platform not only offers a cytocompatible microenvironment for cell encapsulation, but also permits user-controlled rapid recovery of cell structures formed within the hydrogels.

Hydrogels are hydrophilic crosslinked polymers that provide a three-dimensional  microenvironment with tissue-like elasticity and high permeability for  ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

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

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

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

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

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Engineering 'Golden' Fluorescence by Selective Pressure Incorporation of Non-canonical Amino Acids and Protein Analysis by Mass Spectrometry and Fluorescence

Fluorescent proteins are fundamental tools for the life sciences, in particular for fluorescence microscopy of living cells. While wild-type and engineered variants of the green fluorescent protein from Aequorea victoria (avGFP) as well as homologs from other species already cover large parts of the optical spectrum, a spectral gap remains in the near-infrared region, for which avGFP-based fluorophores are not available. Red-shifted fluorescent protein (FP) variants would substantially expand the toolkit for spectral unmixing of multiple molecular species, but the naturally occurring red-shifted FPs derived from corals or sea anemones have lower fluorescence quantum yield and inferior photo-stability compared to the avGFP variants. Further manipulation and possible expansion of the chromophore&#39;s conjugated system towards the far-red spectral region is also limited by the repertoire of 20 canonical amino acids prescribed by the genetic code. To overcome these limitations, synthetic biology can achieve further spectral red-shifting via insertion of non-canonical amino acids into the chromophore triad. We describe the application of SPI to engineer avGFP variants with novel spectral properties. Protein expression is performed in a tryptophan-auxotrophic E. coli strain and by supplementing growth media with suitable indole precursors. Inside the cells, these precursors are converted to the corresponding tryptophan analogs and incorporated into proteins by the ribosomal machinery in response to UGG codons. The replacement of Trp-66 in the enhanced &#34;cyan&#34; variant of avGFP (ECFP) by an electron-donating 4-aminotryptophan results in GdFP featuring a 108 nm Stokes shift and a strongly red-shifted emission maximum (574 nm), while being thermodynamically more stable than its predecessor ECFP. Residue-specific incorporation of the non-canonical amino acid is analyzed by mass spectrometry. The spectroscopic properties of GdFP are characterized by time-resolved fluorescence spectroscopy as one of the valuable applications of genetically encoded FPs in life sciences.

Engineering &#039;Golden&#039; Fluorescence by Selective Pressure Incorporation of Non-canonical Amino Acids and Protein Analysis by Mass Spectrometry and Fluorescence

Synthetic biology enables the engineering of proteins with unprecedented properties using the co-translational insertion of non-canonical amino acids. Here, we presented how a spectrally red-shifted variant of a GFP-type fluorophore with novel fluorescence spectroscopic properties, termed &#34;gold&#34; fluorescent protein (GdFP), is produced in E. coli via selective pressure incorporation (SPI).

Fluorescent proteins are fundamental tools for the life sciences, in particular for fluorescence microscopy of living cells. While wild-type and ...

Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human patients. Human Organ-on-a-Chip (Organ Chip) microfluidic cell culture devices, which provide an experimental in vitro platform to assess efficacy, toxicity, and pharmacokinetic (PK) profiles in humans, may be better predictors of therapeutic efficacy and safety in the clinic compared to animal studies. These devices may be used to model the function of virtually any organ type and can be fluidically linked through common endothelium-lined microchannels to perform in vitro studies on human organ-level and whole body-level physiology without having to conduct experiments on people. These Organ Chips consist of two perfused microfluidic channels separated by a permeable elastomeric membrane with organ-specific parenchymal cells on one side and microvascular endothelium on the other, which can be cyclically stretched to provide organ-specific mechanical cues (e.g., breathing motions in lung). This protocol details the fabrication of flexible, dual channel, Organ Chips through casting of parts using 3D printed molds, enabling combination of multiple casting and post-processing steps. Porous poly (dimethyl siloxane) (PDMS) membranes are cast with micrometer sized through-holes using silicon pillar arrays under compression. Fabrication and assembly of Organ Chips involves equipment and steps that can be implemented outside of a traditional cleanroom. This protocol provides researchers with access to Organ Chip technology for in vitro organ- and body-level studies in drug discovery, safety and efficacy testing, as well as mechanistic studies of fundamental biological processes.

Here, we present a protocol that describes the fabrication of stretchable, dual channel, organ chip microfluidic cell culture devices for recapitulating organ-level functionality in vitro.

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human ...

Conducting Multiple Imaging Modes with One Fluorescence Microscope

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to conventional epi-fluorescence microscopy, various imaging techniques have been developed to achieve specific experimental goals. Some of the widely used techniques include single-molecule fluorescence resonance energy transfer (smFRET), which can report conformational changes and molecular interactions with angstrom resolution, and single-molecule detection-based super-resolution (SR) imaging, which can enhance the spatial resolution approximately ten&#160;to twentyfold compared to diffraction-limited microscopy. Here we present a customer-designed integrated system, which merges multiple imaging methods in one microscope, including conventional epi-fluorescent imaging, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging. Different imaging methods can be achieved easily and reproducibly by switching optical elements. This set-up is easy to adopt by any research laboratory in biological sciences with a need for routine and diverse imaging experiments at a reduced cost and space relative to building separate microscopes for individual purposes.

Here we present a practical guide of building an integrated microscopy system, which merges conventional epi-fluorescent imaging, single-molecule detection-based super-resolution imaging, and multi-color single-molecule detection, including single-molecule fluorescence resonance energy transfer imaging, into one set-up in a cost-efficient way.

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to ...

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate (LAL) Assays

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause inflammation, fever, hypo- or hypertension, and, in extreme cases, can lead to tissue and organ damage that may become fatal. The amounts of endotoxin in pharmaceutical products, therefore, are strictly regulated. Among the methods available for endotoxin detection and quantification, the Limulus Amoebocyte Lysate (LAL) assay is commonly used worldwide. While any pharmaceutical product can interfere with the LAL assay, nano-formulations represent a particular challenge due to their complexity. The purpose of this paper is to provide a practical guide to researchers inexperienced in estimating endotoxins in engineered nanomaterials and nanoparticle-formulated drugs. Herein, practical recommendations for performing three LAL formats including turbidity, chromogenic and gel-clot assays are discussed. These assays can be used to determine endotoxin contamination in nanotechnology-based drug products, vaccines, and adjuvants.

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate LAL Assays

Detection of endotoxins in engineered nanomaterials represents one of the grand challenges in the field of nanomedicine. Here, we present a case study that describes the framework composed of three different LAL formats to estimate potential endotoxin contamination in nanoparticles.