This work was funded by the National Science Foundation (1554417) and National Institutes of Health (DE027769).

1. Preparation of buffers
<ol>
	<li>To prepare the ELISA wash buffer, add 3.94 g Tris-base, 8.77 g NaCl, and 1 g bovine serum albumin (BSA) to 1 L of deionized (DI) water. Add 500 &#181;L polysorbate-20. Adjust the pH to 7.2 using HCl or NaOH.</li>
	<li>To prepare the blocking buffer, add 3.94 g Tris-base, 8.77 g NaCl, and 10 g BSA. Add 500 &#181;L polysorbate-20 to 1 L of DI water. Adjust the pH to 7.2 using HCl or NaOH.</li>
	<li>To prepare the elution buffer (PBS), add 8.01 g NaCl, 2.7 g KCl, 1.42 g Na2HPO4, and 0.24 g KH2PO4 to 1 L DI water. Adjust the pH to 7.4 using HCl or NaOH. 
	&#8203;NOTE: A 10x solution of this buffer can be made and diluted with DI water as needed.</li>
</ol>
2. Preparation of OMV sample
<ol>
	<li>Grow A. actinomycetemcomitans cells to the late exponential phase (optical density at 600 nm of 0.7). Pellet the cells by centrifuging twice at 10,000 x g at 4 &#176;C for 10 min. Filter the supernatant through a 0.45 &#181;m filter.</li>
	<li>Concentrate the bacteria-free supernatant using 50 kDa-molecular weight cut-off filters. Ultracentrifuge the concentrated solution at 105,000 x g at 4 &#176;C for 30 min.&#160;</li>
	<li>Resuspend the pellet in PBS and ultracentrifuge again (105,000 x g at 4 &#176;C for 30 min.) Resuspend the pellet in 2 mL of PBS.</li>
</ol>
3. Packing the S-1000 column
<ol>
	<li>Mix the stock bottle of gel filtration medium with a glass stir rod and pour out into a glass bottle the volume necessary to fill the column, plus approximately 50% excess (about 135 mL). Let these beads sit until they have settled, and then decant off the excess liquid. Resuspend the beads in elution buffer, so that the final solution is approximately 70% (by volume) gel, 30% buffer. Degas the solution under vacuum.</li>
	<li>Mount the glass column vertically using a ring stand and fill with elution buffer to wet the walls of the column. Drain the buffer until there is only about 1 cm of buffer remaining in the column.</li>
	<li>Without creating bubbles, carefully pipette beads into the column, filling the column to the top. Continue to drain excess buffer throughout this process. Be sure to not let the beads settle completely before adding additional beads to the top of the column. The column should be packed to a height of about 2 cm below the bottom of the column reservoir.</li>
</ol>
4. Loading the sample and collecting fractions
<ol>
	<li>Degas the elution buffer under vacuum. Wash the column with two column-volumes (180 mL) of elution buffer.</li>
	<li>Allow the remaining buffer to fully enter the column. Once the buffer has reached the top of the gel layer, carefully pipette a 2-mL sample containing OMVs (at a lipid concentration of approximately 100 - 200 nmol/L) onto the surface of the beads, being careful not to disturb any of the beads at the top of the column. Allow the sample to fully enter the gel, that is, when no liquid remains above the gel layer.&#160;</li>
	<li>Carefully and slowly add elution buffer on top of the gel column. Do not disturb the top layer of the gel, as this will cause sample dilution.</li>
	<li>Place a single 50-mL tube under the column and open the column. Collect the first 20 mL of the eluent. Add additional elution buffer to the top of the column, carefully, as needed to ensure the column is never dry.</li>
	<li>Place a series of 1.5 mL tubes under the column. Start the column and collect a series of 1-mL samples in each tube. As the samples are being collected, continue to add elution buffer to the top of the column, as necessary. Repeat until 96 fractions have been collected. Stop the column. 
	NOTE: The samples should be stored at -20 &#176;C for long-term storage or 4 &#176;C for short-term storage until further analysis.</li>
	<li>To clean the column, run one column volume (90 mL) of 0.1 M NaOH through the column. Run two column volumes (180 mL) of elution buffer through the column.</li>
</ol>
5. Sample analysis
<ol>
	<li>To measure the lipid concentration in each fraction, pipette 50 &#181;L of each fraction into a single well of a 96-well plate. To each well, add 2.5 &#181;L of lipophilic dye. Incubate for 15 s. Measure the fluorescence intensity on a plate reader with an excitation wavelength of 515 nm and an emission wavelength of 640 nm. To calculate the fraction of all lipid in each sample, sum all of the emission intensities and divide each individual intensity by the total.</li>
	<li>To measure the concentration of a particular protein, pipette 100 &#181;L of each fraction into a single well of an ELISA immuno-plate. Incubate at 25 &#176;C for 3 h.
	<ol>
		<li>Decant the samples. Add 200 &#181;L of ELISA wash buffer to each well and decant. Repeat four times for a total of five washes.</li>
		<li>Add 200 &#181;L of blocking buffer to each well and incubate for 1 h at 25 &#176;C. Decant.</li>
		<li>Incubate plates with 100 &#181;L blocking buffer plus primary antibody (1:10,000 for purified antibody; 1:10 for unpurified antibody) overnight at 4 &#176;C. Decant.</li>
		<li>Add 200 &#181;L of ELISA wash buffer to each well and decant. Repeat four times for a total of five washes.</li>
		<li>Add 100 &#181;L of ELISA wash buffer plus secondary antibody (1:30,000) to each well. Incubate for 1 h at 25 &#176;C.</li>
		<li>Add 200 &#181;L of ELISA wash buffer to each well and decant. Repeat four times for a total of five washes.</li>
		<li>Add 100 &#181;L of the 3,3&#39;,5,5&#39;-tetramethylbenzidine (TMB) one-step solution and incubate for 15-30 min or until a blue color develops. Stop the TMB reaction with 50 &#181;L of the stopping solution.</li>
		<li>On a plate reader, read the absorbance of each well at a wavelength of 450 nm.</li>
	</ol>
	</li>
	<li>To measure the total protein concentration, record the absorbance at a wavelength of 280 nm (A280) of each fraction, using a UV-vis spectrophotometer.</li>
</ol>
A schematic of the protocol is shown in Figure 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62429/62429fig01.jpg" /> 
Figure 1: Schematic of SEC procedure. The column is packed with degassed gel filtration medium carefully to avoid bubbles and discontinuities, then washed with two column volumes of elution buffer. Next, the sample is carefully pipetted onto the top of the gel, without disrupting gel packing. The column is opened and run until the sample completely enters the gel. At this point, buffer is placed on the top of the column, and the first 20 mL of eluate is collected. Next, a series of 1-mL fractions is collected. These fractions are then placed in a 96-well plate or 96-well immuno-plate for analysis of lipid and protein content. <a href="https://www.jove.com/files/ftp_upload/62429/62429fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have no conflicts of interest to report.

Here, we have provided a protocol for the simple, fast, and reproducible separation of bacterial OMV subpopulations. Although the technique is relatively straight-forward, there are some steps that must be performed extremely carefully to ensure that efficient separation occurs in the column. First, it is essential that the gel be loaded into the column carefully and slowly to avoid air bubbles. We have observed that leaving the gel at room temperature for several hours before loading the column allows the gel to equilib...

Gram-negative bacteria release vesicles derived from their outer membrane, so-called outer membrane vesicles (OMVs), throughout growth. These OMVs play important roles in cell-to-cell communication, both between bacteria and host as well as between bacterial cells, by carrying a number of important biomolecules, including DNA/RNA, proteins, lipids, and peptidoglycans1,2. In particular, the role of OMVs in bacterial pathogenesis has been extensively studied due to their enrichment in certain virulence factors and toxins3,4,5,6,7,8,9,10,11.
OMVs have been reported to range in size from 20 to 450 nm, depending on the parent bacteria and the growth stage, with several types of bacteria releasing heterogeneously sized OMVs8,12,13,14, which also differ in their protein composition and mechanism of host cell entry12. H. pylori released OMVs ranging in diameter from 20 to 450 nm, with the smaller OMVs containing a more homogeneous protein composition than the larger OMVs. Importantly, the two populations of OMVs were observed to be internalized by host cells via different mechanisms12. In addition, we have demonstrated that Aggregatibacter actinomycetemcomitans releases a population of small (&#60;150 nm) OMVs along with a population of large (&#62;350 nm) OMVs, with the OMVs containing a significant amount of a secreted protein toxin, leukotoxin (LtxA)15. While the role of OMV heterogeneity in cellular processes is clearly important, technical difficulties in separating and analyzing distinct populations of vesicles has limited these studies.
In addition to their importance in bacterial pathogenesis, OMVs have been proposed for use in a number of biotechnological applications, including as vaccines and drug delivery vehicles16,17,18,19,20. For their translational use in such approaches, a clean and monodisperse preparation of vesicles is required. Thus, effective&#160;and efficient methods of separation are necessary.
Most commonly, density gradient centrifugation (DGC) is used to separate heterogeneously sized vesicle populations from cellular debris, including flagellae and secreted proteins21; the method has also been reported as an approach to separate heterogeneously sized OMV subpopulations12,13,14. However, DGC is time-consuming, inefficient, and highly variable user-to-user22 and is, therefore, not ideal for scale-up. In contrast, size exclusion chromatography (SEC) represents a scalable, efficient, and consistent approach to purify OMVs21,23,24. We have found that a long (50-cm), gravity-flow, SEC column, filled with gel filtration medium is sufficient for efficiently purifying and separating subpopulations of OMVs. Specifically, we used this approach to separate A. actinomycetemcomitans OMVs into &#34;large&#34; and &#34;small&#34; subpopulations, as well as to remove protein and DNA contamination. Purification was completed in less than 4 h, and complete separation of the OMV subpopulations and removal of debris was accomplished.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1-Step Ultra TMB-ELISA</td>
			<td>Thermo Scientific</td>
			<td>34028</td>
			<td></td>
		</tr>
		<tr>
			<td>Amicon 50 kDa filters</td>
			<td>Millipore Sigma</td>
			<td>UFC905024</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Fisher Scientific</td>
			<td>BP9704-100</td>
			<td></td>
		</tr>
		<tr>
			<td>ELISA Immuno Plates</td>
			<td>Thermo Scientific</td>
			<td>442404</td>
			<td></td>
		</tr>
		<tr>
			<td>FM 4-64</td>
			<td>Thermo Scientific</td>
			<td>T13320</td>
			<td>1.5 x 50 cm</td>
		</tr>
		<tr>
			<td>Glass Econo-Column</td>
			<td>BioRad</td>
			<td>7371552</td>
			<td></td>
		</tr>
		<tr>
			<td>Infinite 200 Pro Plate Reader</td>
			<td>Tecan</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Chloride (KCl)</td>
			<td>Amresco (VWR)</td>
			<td>0395-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Phosphate Monobasic Anhydrous (KH2PO4)</td>
			<td>Amresco (VWR)</td>
			<td>0781-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sephacryl S-1000 Superfine</td>
			<td>GE Healthcare</td>
			<td>17-0476-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride (NaCl)</td>
			<td>Fisher Chemical</td>
			<td>S271-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Phosphate Dibasic Anhydrous (Na2HPO4)</td>
			<td>Amresco (VWR)</td>
			<td>0404-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris Base</td>
			<td>VWR</td>
			<td>0497-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween(R) 20</td>
			<td>Acros Organics</td>
			<td>23336-2500</td>
			<td></td>
		</tr>
	</tbody>
</table>

size exclusion chromatography to analyze bacterial outer membrane vesicle heterogeneity

The cell wall of Gram-negative bacteria consists of an inner (cytoplasmic) and outer membrane (OM), separated by a thin peptidoglycan layer. Throughout growth, the outer membrane can bleb to form spherical outer membrane vesicles (OMVs). These OMVs are involved in numerous cellular functions including cargo delivery to host cells and communication with bacterial cells. Recently, the therapeutic potential of OMVs has begun to be explored, including their use as vaccines and drug delivery vehicles. Although OMVs are derived from the OM, it has long been appreciated that the lipid and protein cargo of the OMV differs, often significantly, from that of the OM. More recently, evidence that bacteria can release multiple types of OMVs has been discovered, and evidence exists that size can impact the mechanism of their uptake by host cells. However, studies in this area are limited by difficulties in efficiently separating the heterogeneously sized OMVs. Density gradient centrifugation (DGC) has traditionally been used for this purpose; however, this technique is time-consuming and difficult to scale-up. Size exclusion chromatography (SEC), on the other hand, is less cumbersome and lends itself to the necessary future scale-up for therapeutic use of OMVs. Here, we describe a SEC approach that enables reproducible separation of heterogeneously sized vesicles, using as a test case, OMVs produced by Aggregatibacter actinomycetemcomitans, which range in diameter from less than 150 nm to greater than 350 nm. We demonstrate separation of &#34;large&#34; (350 nm) OMVs and &#34;small&#34; (&#60;150 nm) OMVs, verified by dynamic light scattering (DLS). We recommend SEC-based techniques over DGC-based techniques for separation of heterogeneously sized vesicles due to its ease of use, reproducibility (including user-to-user), and possibility for scale-up.

Figure 2&#160;shows representative results from this method. OMVs produced by A. actinomycetemcomitans strain JP2 were first purified from the culture supernatant using ultracentrifugation15. We previously found that this strain produces two populations of OMVs, one with diameters of about 300 nm and one with diameters of about 100 nm15. To separate these OMV populations, we purified the sample using the SEC protocol described above. E...

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Size Exclusion Chromatography to Analyze Bacterial Outer Membrane Vesicle Heterogeneity

Bacterial vesicles play important roles in pathogenesis and have promising biotechnological applications. The heterogeneity of vesicles complicates analysis and use; therefore, a simple, reproducible method to separate varying sizes of vesicles is necessary. Here, we demonstrate the use of size exclusion chromatography to separate heterogeneous vesicles produced by Aggregatibacter actinomycetemcomitans.

The cell wall of Gram-negative bacteria consists of an inner (cytoplasmic) and outer membrane (OM), separated by a thin peptidoglycan layer. Throughout ...

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Bioengineering

4.2K Views.  Lehigh University. Bacterial vesicles play important roles in pathogenesis and have promising biotechnological applications. The heterogeneity of vesicles complicates analysis and use; therefore, a simple, reproducible method to separate varying sizes of vesicles is necessary. Here, we demonstrate the use of size exclusion chromatography to separate heterogeneous vesicles produced by Aggregatibacter actinomycetemcomitans.

Video: Size Exclusion Chromatography to Analyze Bacterial Outer Membrane Vesicle Heterogeneity

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Bacterial Detection &amp; Identification Using Electrochemical Sensors

Electrochemical sensors are widely used for rapid and accurate measurement of blood glucose and can be adapted for detection of a wide variety of analytes. Electrochemical sensors operate by transducing a biological recognition event into a useful electrical signal. Signal transduction occurs by coupling the activity of a redox enzyme to an amperometric electrode. Sensor specificity is either an inherent characteristic of the enzyme, glucose oxidase in the case of a glucose sensor, or a product of linkage between the enzyme and an antibody or probe.
Here, we describe an electrochemical sensor assay method to directly detect and identify bacteria. In every case, the probes described here are DNA oligonucleotides. This method is based on sandwich hybridization of capture and detector probes with target ribosomal RNA (rRNA). The capture probe is anchored to the sensor surface, while the detector probe is linked to horseradish peroxidase (HRP). When a substrate such as 3,3',5,5'-tetramethylbenzidine (TMB) is added to an electrode with capture-target-detector complexes bound to its surface, the substrate is oxidized by HRP and reduced by the working electrode. This redox cycle results in shuttling of electrons by the substrate from the electrode to HRP, producing current flow in the electrode.

Bacterial Detection & Identification Using Electrochemical Sensors

We describe an electrochemical sensor assay method for rapid bacterial detection and identification. The assay involves a sensor array functionalized with DNA oligonucleotide capture probes for ribosomal RNA (rRNA) species-specific sequences. Sandwich hybridization of target rRNA with the capture probe and a horseradish peroxidase-linked DNA oligonucleotide detector probe produces a measurable amperometric current.

Electrochemical  sensors are widely used for rapid and accurate measurement of blood glucose and  can be adapted for detection of a wide variety of ...

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to identify and characterize their interactions are necessary5. To this end, we developed the Membrane Strep-protein interaction experiment, called Membrane-SPINE6. This technique combines in vivo cross-linking using the reversible cross-linker formaldehyde with affinity purification of a Strep-tagged membrane bait protein. During the procedure, cross-linked prey proteins are co-purified with the membrane bait protein and subsequently separated by boiling. Hence, two major tasks can be executed when analyzing protein-protein interactions (PPIs) of membrane proteins using Membrane-SPINE: first, the confirmation of a proposed interaction partner by immunoblotting, and second, the identification of new interaction partners by mass spectrometry analysis. Moreover, even low affinity, transient PPIs are detectable by this technique. Finally, Membrane-SPINE is adaptable to almost any cell type, making it applicable as a powerful screening tool to identify PPIs of membrane proteins.

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

A biochemical approach is described to identify in vivo protein-protein interactions (PPI) of membrane proteins. The method combines protein cross-linking, affinity purification and mass spectrometry, and is adaptable to almost any cell type or organism. With this approach, even the identification of transient PPIs becomes possible.

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to ...

Microfluidic Fabrication of Polymeric and Biohybrid Fibers with Predesigned Size and Shape

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and used to dictate the shape as well as the diameter of a core stream.&#160;Grooves in the top and bottom of a microfluidic channel were designed to direct the sheath fluid and shape the core fluid.&#160;By matching the viscosity and hydrophilicity of the sheath and core fluids, the interfacial effects are minimized and complex fluid shapes can be formed.&#160;Controlling the relative flow rates of the sheath and core fluids determines the cross-sectional area of the core fluid.&#160;Fibers have been produced with sizes ranging from 300 nm to ~1 mm, and fiber cross-sections can be round, flat, square, or complex as in the case with double anchor fibers. Polymerization of the core fluid downstream from the shaping region solidifies the fibers.&#160;Photoinitiated click chemistries are well suited for rapid polymerization of the core fluid by irradiation with ultraviolet light.&#160;Fibers with a wide variety of shapes have been produced from a list of polymers including liquid crystals, poly(methylmethacrylate), thiol-ene and thiol-yne resins, polyethylene glycol, and hydrogel derivatives. Minimal shear during the shaping process and mild polymerization conditions also makes the fabrication process well suited for encapsulation of cells and other biological components.

Two adjacent fluids passing through a grooved microfluidic channel can be directed to form a sheath around a prepolymer core; thereby&#160;determining both shape and cross-section. Photoinitiated polymerization, such as thiol click chemistry, is well suited for rapidly solidifying the core fluid into a microfiber with predetermined size and shape.

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and ...

Lipid Bilayer Vesicle Generation Using Microfluidic Jetting

Bottom-up synthetic biology presents a novel approach for investigating and reconstituting biochemical systems and, potentially, minimal organisms. This emerging field engages engineers, chemists, biologists, and physicists to design and assemble basic biological components into complex, functioning systems from the bottom up. Such bottom-up systems could lead to the development of artificial cells for fundamental biological inquiries and innovative therapies1,2. Giant unilamellar vesicles (GUVs) can serve as a model platform for synthetic biology due to their cell-like membrane structure and size. Microfluidic jetting, or microjetting, is a technique that allows for the generation of GUVs with controlled size, membrane composition, transmembrane protein incorporation, and encapsulation3. The basic principle of this method is the use of multiple, high-frequency fluid pulses generated by a piezo-actuated inkjet device to deform a suspended lipid bilayer into a GUV. The process is akin to blowing soap bubbles from a soap film. By varying the composition of the jetted solution, the composition of the encompassing solution, and/or the components included in the bilayer, researchers can apply this technique to create customized vesicles. This paper describes the procedure to generate simple vesicles from a droplet interface bilayer by microjetting.

Microfluidic jetting against a droplet interface lipid bilayer provides a reliable way to generate vesicles with control over membrane asymmetry, incorporation of transmembrane proteins, and encapsulation of material. This technique can be applied to study a variety of biological systems where compartmentalized biomolecules are desired.

Bottom-up synthetic biology presents a novel approach for investigating and reconstituting biochemical systems and, potentially, minimal organisms. This ...

Quantification and Size-profiling of Extracellular Vesicles Using Tunable Resistive Pulse Sensing

Extracellular vesicles (EVs), including &#8216;microvesicles&#8217; and &#8216;exosomes&#8217;, are highly abundant in bodily fluids. Recent years have witnessed a tremendous increase in interest in EVs. EVs have been shown to play important roles in various physiological and pathological processes, including coagulation, immune responses, and cancer. In addition, EVs have potential as therapeutic agents, for instance as drug delivery vehicles or as regenerative medicine. Because of their small size (50 to 1,000 nm) accurate quantification and size profiling of EVs is technically challenging.
This protocol describes how tunable resistive pulse sensing (tRPS) technology, using the qNano system, can be used to determine the concentration and size of EVs. The method, which relies on the detection of EVs upon their transfer through a nano sized pore, is relatively fast, suffices the use of small sample volumes and does not require the purification and concentration of EVs. Next to the regular operation protocol an alternative approach is described using samples spiked with polystyrene beads of known size and concentration. This real-time calibration technique can be used to overcome technical hurdles encountered when measuring EVs directly in biological fluids.

Extracellular vesicles play important roles in physiological and pathological processes, including coagulation, immune responses, and cancer or as potential therapeutic agents in drug delivery or regenerative medicine. This protocol presents methods for the quantification and size characterization of isolated and non-isolated extracellular vesicles in various fluids using tunable resistive pulse sensing.

Extracellular vesicles (EVs), including &#8216;microvesicles&#8217; and &#8216;exosomes&#8217;, are highly abundant in bodily fluids. Recent years have ...

Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range of 10-100 &#197;. By measuring the distances under various ligated conditions, conformational changes of the protein can be easily assessed. With LRET, a lanthanide, most often chelated terbium, is used as the donor fluorophore, affording advantages such as a longer donor-only emission lifetime, the flexibility to use multiple acceptor fluorophores, and the opportunity to detect sensitized acceptor emission as an easy way to measure energy transfer without the risk of also detecting donor-only signal. Here, we describe a method to use LRET on membrane proteins expressed and assayed on the surface of intact mammalian cells. We introduce a protease cleavage site between the LRET fluorophore pair. After obtaining the original LRET signal, cleavage at that site removes the specific LRET signal from the protein of interest allowing us to quantitatively subtract the background signal that remains after cleavage. This method allows for more physiologically relevant measurements to be made without the need for purification of protein.

We describe here an improved Luminescence Resonance Energy Transfer (LRET) method where we introduce a protease cleavage site between the donor and acceptor fluorophore sites. This modification allows us to obtain specific LRET signals arising from membrane proteins of interest, allowing for the study of membrane proteins without protein purification.

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range ...

Methods for Characterizing the Co-development of Biofilm and Habitat Heterogeneity

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is strongly regulated by the surrounding flow and nutritional environment. Biofilm growth also increases the heterogeneity of the local microenvironment by generating complex flow fields and solute transport patterns. To investigate the development of heterogeneity in biofilms and interactions between biofilms and their local micro-habitat, we grew mono-species biofilms of Pseudomonas aeruginosa and dual-species biofilms of P. aeruginosa and Escherichia coli under nutritional gradients in a microfluidic flow cell. We provide detailed protocols for creating nutrient gradients within the flow cell and for growing and visualizing biofilm development under these conditions. We also present protocols for a series of optical methods to quantify spatial patterns in biofilm structure, flow distributions over biofilms, and mass transport around and within biofilm colonies. These methods support comprehensive investigations of the co-development of biofilm and habitat heterogeneity.

Biofilms have complex interactions with their surrounding environment. To comprehensively investigate biofilm-environment interactions, we present here a series of methods to create heterogeneous chemical environment for biofilm development, to quantify local flow velocity, and to analyze mass transport in and around biofilm colonies.

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is ...

A Facile and Eco-friendly Route to Fabricate Poly(Lactic Acid) Scaffolds with Graded Pore Size

Over the recent years, functionally graded scaffolds (FGS) gaineda crucial role for manufacturing of devices for tissue engineering. The importance of this new field of biomaterials research is due to the necessity to develop implants capable of mimicking the complex functionality of the various tissues, including a continuous change from one structure or composition to another. In this latter context, one topic of main interest concerns the design of appropriate scaffolds for bone-cartilage interface tissue. In this study, three-layered scaffolds with graded pore size were achieved by melt mixing poly(lactic acid) (PLA), sodium chloride (NaCl) and polyethylene glycol (PEG). Pore size distributions were controlled by NaCl granulometry and PEG solvation. Scaffolds were characterized from a morphological and mechanical point of view. A correlation between the preparation method, the pore architecture and compressive mechanical behavior was found. The interface adhesion strength was quantitatively evaluated by using a custom-designed interfacial strength test. Furthermore, in order to imitate the human physiology, mechanical tests were also performed in phosphate buffered saline (PBS) solution at 37 &#176;C. The method herein presented provides a high control of porosity, pore size distribution and mechanical performance, thus offering the possibility to fabricate three-layered scaffolds with tailored properties by following a simple and eco-friendly route.

A Facile and Eco-friendly Route to Fabricate PolyLactic Acid Scaffolds with Graded Pore Size

In this work, poly(lactic acid)/polyethylene glycol (PLA/PEG) scaffolds were prepared by using a combination of melt mixing and selective leaching. The method herein discussed permitted to develop three-layer scaffolds by highly controlling both porosity and pore size. The mechanical properties were also evaluated in a physiological environment.

Over the recent years, functionally graded scaffolds (FGS) gaineda crucial role for manufacturing of devices for tissue engineering. The importance of ...

Treatment of Ligament Constructs with Exercise-conditioned Serum: A Translational Tissue Engineering Model

In vitro experiments are essential to understand biological mechanisms; however, the gap between monolayer tissue culture and human physiology is large, and translation of findings is often poor. Thus, there is ample opportunity for alternative experimental approaches. Here we present an approach in which human cells are isolated from human anterior cruciate ligament tissue remnants, expanded in culture, and used to form engineered ligaments. Exercise alters the biochemical milieu in the blood such that the function of many tissues, organs and bodily processes are improved. In this experiment, ligament construct culture media was supplemented with experimental human serum that has been &#39;conditioned&#39; by exercise. Thus the intervention is more biologically relevant since an experimental tissue is exposed to the full endogenous biochemical milieu, including binding proteins and adjunct compounds that may be altered in tandem with the activity of an unknown agent of interest. After treatment, engineered ligaments can be analyzed for mechanical function, collagen content, morphology, and cellular biochemistry. Overall, there are four major advantages versus traditional monolayer culture and animal models, of the physiological model of ligament tissue that is presented here. First, ligament constructs are three-dimensional, allowing for mechanical properties (i.e., function) such as ultimate tensile stress, maximal tensile load, and modulus, to be quantified. Second, the enthesis, the interface between boney and sinew elements, can be examined in detail and within functional context. Third, preparing media with post-exercise serum allows for the effects of the exercise-induced biochemical milieu, which is responsible for the wide range of health benefits of exercise, to be investigated in an unbiased manner. Finally, this experimental model advances scientific research in a humane and ethical manner by replacing the use of animals, a core mandate of the National Institutes of Health, the Center for Disease Control, and the Food and Drug Administration.

We present a model of ligament tissue in which three-dimensional constructs are treated with the human exercise-conditioned serum and analyzed for collagen content, function, and cellular biochemistry.

In vitro experiments are essential to understand biological mechanisms; however, the gap between monolayer tissue culture and human physiology is large, ...