This research was supported by grants of the Deutsche Forschungsgemeinschaft GraKo1121, Hu1011/2-1 and SFB940 to S.H.

Note: Detailed information regarding buffers indicated in the protocol is available in Table 1.
1. Fixation of Protein-protein Interactions by Formaldehyde Cross-linking in Living Cells
<ol>
	<li>To begin prepare growth media, stock solutions and buffer P1
	<ol>
		<li>Prepare 500 ml medium: The medium should provide conditions that support the interaction between membrane bait protein and prey proteins. In addition, one experiment should be performed under conditions where no interaction is expected (e.g. without cross-linking). 
		Note: We compare protein-protein interactions in stressed and non-stressed bacteria. Therefore, we prepare 500 ml Luria Bertani (LB) medium (Table 1) that we complement with a stress-inducing agent (e.g. 0.5 M NaCl) in a 2 L Erlenmeyer flask. For control, we use normal LB medium with 0.17 M NaCl (pH 7.0).</li>
		<li>Prepare Tris-buffer (20 mM Tris-HCl; pH 8.0) and 0.1 M ethylenediaminetetraacetic acid (EDTA), pH 8.0 stock solution (Table 1).</li>
		<li>Prepare buffer P1. Dissolve sucrose to a final concentration of 0.5 M in Tris-buffer. Sterilize buffer P1 by filtration and store it at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Grow cells expressing the membrane bait protein with a C-terminal Strep-tag fusion from a vector. Induce the expression of membrane bait proteins for a sufficient time. 
	Note: We induce the expression of bacterial membrane bait proteins at early-log phase (A600 = 0.3) by addition of 250 &#181;l 1 M isopropyl-&#946;-D-thiogalactopyranoside (IPTG) to 500 ml medium (final concentration: 0.5 mM). To allow sufficient expression, we incubate the bacteria until the late-log phase (A600 = 1.2).</li>
	<li>Prepare for each culture two centrifuge beakers with a minimum volume of 300 ml each and place them on ice.</li>
	<li>Perform formaldehyde cross-linking. 
	CAUTION: Formaldehyde is highly toxic. We recommend working under a safety fume hood.
	<ol>
		<li>Transfer the culture vessel under a safety fume hood. Split each sample into two samples, one omitting cross-linking (- X) and one including the formaldehyde cross-linking (+ X). Add 4 ml 37% formaldehyde solution to 250 ml culture to reach a final concentration of 0.6%.</li>
	</ol>
	</li>
	<li>Transfer the culture vessels back and grow cells for further 20 min as before.</li>
	<li>Fill your cultures into the centrifuge tubes under a safety fume hood. Collect cells by centrifugation at 3,000 x g for 30 min.</li>
	<li>Discard supernatant by carefully pipetting it up under a safety fume hood. Collect toxic formaldehyde-containing supernatant and dispose of it properly.</li>
	<li>In order to reach optimal yield during membrane protein preparation, prepare spheroplasts first. Here, we provide a protocol for spheroplast formation of Gram-negative bacteria:
	<ol>
		<li>Prepare buffer P2 (Table 1) from lyophilized powder. Gently dissolve 2 mg lysozyme in 0.1 M EDTA, pH 8, in a 1.5 ml tube. Always prepare buffer P2 freshly for the day of the experiment.</li>
		<li>Prepare Tris-buffer with protease inhibitor (buffer P3). To 10 ml of Tris-buffer, add 0.1 ml 1 M phenylmethylsulfonylfluoride (PMSF) (Table 1) to a final concentration of 10 mM. Use the buffer immediately, to ensure proper function of PMSF as protease inhibitor.</li>
		<li>Resuspend cell pellets in 10 ml Tris-buffer with protease inhibitor and transfer solution to a 15 ml conical tube.</li>
		<li>Add 1 ml buffer P2 and incubate on ice for 30 min.</li>
		<li>Collect spheroplasts by centrifugation at 3,000 x g for 30 min and discard supernatant carefully.</li>
	</ol>
	</li>
	<li>Incubate the spheroplast pellets overnight at -20&#176;C.</li>
</ol>
2. Purification of Strep-tag Membrane Protein (Bait)
<ol>
	<li>Place spheroplast pellet on ice.</li>
	<li>During the time the spheroplast pellets thaw on ice, prepare 10 ml buffer P3 for each spheroplast preparation. Gently dissolve 1 mg DNaseI in 10 ml Tris-buffer. Add 0.1 ml 1 M PMSF just before use.</li>
	<li>Resuspend the pellet in 6 ml freshly prepared P3. Sonicate the sample four times for 1 min continuously on ice with 1 min pause between each burst, in order to disrupt spheroplasts.</li>
	<li>Centrifuge the sample at 10,000 x g for 10 min to harvest cell debris. Transfer the supernatant using a pipette to an ultracentrifuge tube.</li>
	<li>Pellet the membrane fraction at 100,000 x g for 30 min. Wash the pellet carefully in Tris-buffer without dissolving it. Dry the tube with a paper tissue (e.g. Kleenex). Avoid disturbing the pellet at any time.</li>
	<li>Resuspend the pellet (= membranes) carefully in 1 ml Tris-buffer. Use a 20 &#181;l aliquot to determine the protein concentration by e.g. BCA assay. At this step, the membranes can be shock frozen in liquid nitrogen and stored at -80 &#176;C.</li>
</ol>
3. Purification of Strep-tag Membrane Bait Protein and SDS-PAGE
<ol>
	<li>Normalize the protein concentration of the membrane fraction to 5 mg/ml with Tris-buffer. Take 2.5 ml membrane fraction (5 mg/ml) in an ultracentrifuge tube and add 0.25 ml of 20% Triton X-100 to reach a final concentration of 2%, in order to solubilize the membrane proteins. Add a micro-magnetic rod and stir on ice for 1 hr. 
	Note: Although in general we have good results using Triton X-100 as detergent, it might be useful to change the detergent for optimization. In this respect, we have best results with those detergents used for functional purification of the respective membrane bait protein.</li>
	<li>While performing step 3.1, prepare 50 ml buffer W and 5 ml buffer E from (Table 1). Fill up 10 ml 5x buffer W concentrate to 50 ml and add 150 &#181;l 20% Triton X-100.
	<ol>
		<li>Fill up 1 ml 5x buffer E concentrate to 10 ml and add 30 &#181;l 20% Triton X-100. Equilibrate a 1 ml Strep-Tactin superflow gravity flow column with 8 ml buffer W. 
		Note: It is essential&#160;to add the detergent used for solubilization in a concentration 10-fold above the critical micelle concentration (cmc) in any buffer during the purification procedure.</li>
	</ol>
	</li>
	<li>Remove the micro-magnetic rod from the solubilization sample and ultracentrifuge at 100,000 x g for 30 min, to pellet the insoluble membrane fraction. Remove the supernatant using a pipette for further purification.</li>
	<li>Load the supernatant to the column. Run the column only with gravity flow. Wash the column with 5 ml buffer W. Repeat this washing step 5x.</li>
	<li>Elute membrane bait proteins with 1 ml buffer E. Repeat this elution step 4x.</li>
	<li>Concentrate elution fractions 2, 3, and 4 to 300 &#181;l with a centrifugal filter unit.</li>
	<li>Mix 200 &#181;l of each sample with 50 &#181;l 5x SDS-PAGE loading dye. Split each preparation in 125 &#181;l aliquots. Boil one aliquot of each preparation for 20 min at 95 &#176;C, to reverse formaldehyde cross-links. Let samples cool down to RT for at least 10 min on bench top.</li>
	<li>Load 30 &#181;l from each sample to a single lane of a polyacrylamide-gel suitable for immunoblotting. Use a prestained molecular weight marker for best orientation and run SDS-PAGE10.</li>
</ol>
4. Immunoblot Analysis to Confirm Interaction Partners
<ol>
	<li>Once step 3.8 is completed, transfer proteins to a nitrocellulose membrane by e.g. semidry.</li>
	<li>Block the membrane to prevent unspecific labeling. Prepare blocking buffer by weighing bovine serum albumin (BSA) to achieve 3% weight per volume in Tris-buffered saline supplemented with final a concentration of 0.05% Tween-20 (TBS-T). Use a sufficient volume of blocking buffer to cover the membrane. Block the membrane for 1 hr at RT.</li>
	<li>Dilute and incubate the prey protein specific first antibody as usual. Use an HRP-linked antibody as the second antibody.</li>
	<li>Develop the immunoblot using a chemiluminescent detection kit with high sensitivity. Monitor the signal using a classical film processing procedure or digital imaging equipment.</li>
</ol>
5. NanoLC-ESI-MS/MS High Resolution Experiments to Identify Interaction Partners
<ol>
	<li>In case that no specific antibody is available for the prey protein or unknown interaction partners should be identified, use mass spectrometry (MS) for identification. In order to prevent keratin contamination, use precasted gels for protein separation.</li>
	<li>Silver stain the SDS-PAGE using a MS-compatible staining kit according to the manufacturer&#39;s protocol. Perform all staining and washing steps in glass tanks.</li>
	<li>Excise respective bands and analyze these by high resolution LC/MS9.</li>
</ol>

<ol>
	<li>Krogh, A., Larsson, B., von Heijne, G., Sonnhammer, E. L. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Predicting+transmembrane+protein+topology+with+a+hidden+markov+model:+application+to+complete+genomes.">Predicting transmembrane protein topology with a hidden markov model: application to complete genomes.</a> J. Mol. Biol. 305, 567-580 (2001).</li><li>Bakheet, T. M., Doig, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Properties+and+identification+of+human+protein+drug+targets.">Properties and identification of human protein drug targets.</a> Bioinformatics. 25, 451-457 (2009).</li><li>Yildirim, M. A., Goh, K. I., Cusick, M. E., Barabasi, A. L., Vidal, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drug-target+network.">Drug-target network.</a> Nat. Biotechnol. 25, (2007).</li><li>Daley, D. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+assembly+of+membrane+proteins+into+complexes.">The assembly of membrane proteins into complexes.</a> Curr. Opin. Struct. Biol. 18, 420-424 (2008).</li><li>Jung, K., Fried, L., Behr, S., Heermann, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Histidine+kinases+and+response+regulators+in+networks.">Histidine kinases and response regulators in networks.</a> Curr. Opin. Microbiol. 15, 118-124 (2012).</li><li>M&#252;ller, V. S., Jungblut, P. R., Meyer, T. F., Hunke, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane-SPINE:+An+improved+method+to+identify+protein-protein+interaction+partners+of+membrane+proteins+in+vivo.">Membrane-SPINE: An improved method to identify protein-protein interaction partners of membrane proteins in vivo.</a> Proteomics. 11, 2124-2128 (2011).</li><li>Herzberg, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SPINE:+A+method+for+the+rapid+detection+and+analysis+of+protein-protein+interactions+in+vivo.">SPINE: A method for the rapid detection and analysis of protein-protein interactions in vivo.</a> Proteomics. 7, 4032-4035 (2007).</li><li>Sutherland, B. W., Toews, J., Kast, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Utility+of+formaldehyde+cross-linking+and+mass+spectrometry+in+the+study+of+protein-protein+interactions.">Utility of formaldehyde cross-linking and mass spectrometry in the study of protein-protein interactions.</a> J. Mass Spectrom. 43, 699-715 (2008).</li><li>Hernandez, P., M&#252;ller, M., Appel, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+protein+identification+by+tandem+mass+spectrometry:+Issues+and+strategies.">Automated protein identification by tandem mass spectrometry: Issues and strategies.</a> Mass Spectrom. Rev. 25, 235-254 (2006).</li><li>Laemmli, U. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cleavage+of+structural+proteins+during+the+assembly+of+the+head+of+bacteriophage+T4.">Cleavage of structural proteins during the assembly of the head of bacteriophage T4.</a> Nature. 227, 680-685 (1970).</li><li>Hunke, S., Keller, R., M&#252;ller, V. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signal+integration+by+the+Cpx-envelope+stress+system.">Signal integration by the Cpx-envelope stress system.</a> FEMS Microbiol. Lett. 326, 12-22 (2012).</li><li>Weber, R. F., Silverman, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Cpx+proteins+of+Escherichia+coli+K12+:+Structure+of+the+CpxA+polypeptide+as+an+inner+membrane+component.">The Cpx proteins of Escherichia coli K12 : Structure of the CpxA polypeptide as an inner membrane component.</a> J. Mol. Biol. 203, 467-478 (1988).</li></ol>

We have nothing to disclose.

Membrane SPINE analysis is a biochemical approach that enables one to confirm and to identify to this point unknown interaction partners of membrane proteins. Membrane SPINE combines in vivo cross-linking by formaldehyde with purification of a Strep-tagged membrane bait protein. The combination with immunoblotting facilitates the confirmation of predicted interaction partners (Figure 2). Additionally, the combination with MS analysis permits the identification of unknown interaction partners (<s...

To understand the function of a protein it is essential to know its interaction partners. Several classical techniques are available for the identification of interaction partners of soluble proteins. However, these techniques are not easily transferable to membrane proteins due to their hydrophobic nature4. To overcome this limitation, we have developed the Membrane Strep-protein interaction experiment (Membrane-SPINE)6. It is based on the SPINE method, which was only suitable for soluble proteins7.
Membrane-SPINE benefits from two advantages of the cross-linking agent formaldehyde: first, formaldehyde can easily penetrate membranes and therefore generates a precise snapshot of the interactome of a living cell8. Second, formaldehyde cross-links can be reversed by boiling9. Here, these two advantages are used to identify not only permanent but also transient PPIs of membrane proteins6.
In brief, a Strep-tag is fused to the C-terminus of the integral membrane bait protein. Cells expressing the membrane bait protein are incubated with formaldehyde which cross-links prey proteins to the membrane bait protein (Figure 1). Modifications of prey proteins are not needed. Next, the membrane fraction is prepared. Therefore, membrane proteins are solubilized by detergent treatment and bait proteins are co-purified with its prey proteins using affinity purification. Subsequently, the cross-link is reversed by boiling, and the bait and its co-eluted prey proteins are separated by SDS-PAGE. Finally, prey proteins can be identified by immunoblot analysis or mass spectrometry.

<table border="0" cellpadding="0" cellspacing="0" width="878">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">37% Formaldehyde</td>
			<td style="width:128px;">Roth</td>
			<td style="width:137px;">4979.1</td>
			<td style="width:271px;">Should not be older than one year</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">DNaseI</td>
			<td style="width:128px;">Sigma</td>
			<td style="width:137px;">DN25</td>
			<td style="width:271px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">BCA protein assay kit</td>
			<td style="width:128px;">Pierce</td>
			<td style="width:137px;">23225</td>
			<td style="width:271px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Micromagnetic rod</td>
			<td style="width:128px;">Roth</td>
			<td style="width:137px;">0955.2</td>
			<td style="width:271px;">5 mm in length, 2 mm in diameter</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Triton X-100</td>
			<td style="width:128px;">Roth</td>
			<td style="width:137px;">6683.1</td>
			<td style="width:271px;">standard detergent for solubilization</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">n-Dodecyl-&#946;-maltoside</td>
			<td style="width:128px;">Glycon</td>
			<td style="width:137px;">D97002-C</td>
			<td style="width:271px;">the best detergent to solubilze CpxA</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">1 ml Strep-Tactin superflow gravity flow column</td>
			<td style="width:128px;">IBA</td>
			<td style="width:137px;">2-1207-050</td>
			<td style="width:271px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Strep-tag protein purification buffer set</td>
			<td style="width:128px;">IBA</td>
			<td style="width:137px;">2-1002-001</td>
			<td style="width:271px;">Contains buffer W and buffer E</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Amicon Ultra-4 centrifugal filter</td>
			<td style="width:128px;">Millipore</td>
			<td style="width:137px;">UFC803024</td>
			<td style="width:271px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:343px;">SuperSignal West Pico Chemiluminescent substrate</td>
			<td style="width:128px;">Pierce</td>
			<td style="width:137px;">34080</td>
			<td style="width:271px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">SilverQuest Silverstaining kit</td>
			<td style="width:128px;">Invitrogen</td>
			<td style="width:137px;">LC6070</td>
			<td style="width:271px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">FireSilver Staining Kit</td>
			<td style="width:128px;">Proteome Factory</td>
			<td style="width:137px;">P-S-2001</td>
			<td style="width:271px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Ultracentrifuge</td>
			<td style="width:128px;"></td>
			<td style="width:137px;"></td>
			<td style="width:271px;"></td>
		</tr>
	</tbody>
</table>

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 analysis allows the co-purification of membrane proteins and transiently interacting protein partners. The co-purification is achieved by using the cross-linking agent formaldehyde. Two parameters are critical to prevent unspecific cross-links: the formaldehyde concentration and the cross-linking time. The sufficient, but not excessive use of formaldehyde can easily be controlled by immunoblotting. Formaldehyde cross-linked protein complexes can be separated by boiling but not by SDS treatment. Hence, they...

Watch this Scientific Journal Video about Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo at JoVE.com

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-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 ...

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13.6K Views.  Universität Osnabrück. 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.

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

Fabrication of Electrochemical-DNA Biosensors for the Reagentless Detection of Nucleic Acids, Proteins and Small Molecules

As medicine is currently practiced, doctors send specimens to a central laboratory for testing and thus must wait hours or days to 
receive the results. Many patients would be better served by rapid, bedside tests. To this end our laboratory and others have developed a versatile, reagentless 
biosensor platform that supports the quantitative, reagentless, electrochemical detection of nucleic acids (DNA, RNA), proteins (including antibodies) and small 
molecules analytes directly in unprocessed clinical and environmental samples. In this video, we demonstrate the preparation and use of several biosensors in this 
"E-DNA" class. In particular, we fabricate and demonstrate sensors for the detection of a target DNA sequence in a polymerase chain reaction mixture, an HIV-specific antibody and the drug cocaine. The preparation procedure requires only three hours of hands-on effort followed by an overnight incubation, and their use requires only minutes.

"E-DNA" sensors, reagentless, electrochemical biosensors that perform well even when challenged directly in blood and other complex matrices, have been adapted to the detection of a wide range of nucleic acid, protein and small molecule analytes. Here we present a general procedure for the fabrication and use of such sensors.

As medicine is currently practiced, doctors send specimens to a central laboratory for testing and thus must wait hours or days to 
receive the results. ...

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural biology 1. In recent years, this method has achieved great success in studying structures of proteins and macromolecular complexes 2, 3. Compared with electron cryomicroscopy (cryoEM), in which frozen hydrated protein samples are embedded in a thin layer of vitreous ice 4, negative staining is a simpler sample preparation method in which protein samples are embedded in a thin layer of dried heavy metal salt to increase specimen contrast 5. The enhanced contrast of negative stain EM allows examination of relatively small biological samples. In addition to determining three-dimensional (3D) structure of purified proteins or protein complexes 6, this method can be used for much broader purposes. For example, negative stain EM can be easily used to visualize purified protein samples, obtaining information such as homogeneity/heterogeneity of the sample, formation of protein complexes or large assemblies, or simply to evaluate the quality of a protein preparation.
In this video article, we present a complete protocol for using an EM to observe negatively stained protein sample, from preparing carbon coated grids for negative stain EM to acquiring images of negatively stained sample in an electron microscope operated at 120kV accelerating voltage. These protocols have been used in our laboratory routinely and can be easily followed by novice users.

Visualizing protein samples by negative stain electron microscopy (EM) has become a popular structural analysis method. It is useful for quantitative structural analysis, such as calculating a 3D reconstruction of the molecules being studied, and also for qualitative examination of the quality of protein preparations. In this article we present detailed protocols for preparing the EM grids, staining the sample and visualizing the sample in an electron microscope. Novice users can follow these protocols easily and to utilize negative stain EM as a routine assay, in addition to other biochemical assays, for evaluating their protein samples.

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural ...

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 ...

Determination of the Transport Rate of Xenobiotics and Nanomaterials Across the Placenta using the ex vivo Human Placental Perfusion Model

Decades ago the human placenta was thought to be an impenetrable barrier between mother and unborn child. However, the discovery of thalidomide-induced birth defects and many later studies afterwards proved the opposite. Today several harmful xenobiotics like nicotine, heroin, methadone or drugs as well as environmental pollutants were described to overcome this barrier. With the growing use of nanotechnology, the placenta is likely to come into contact with novel nanoparticles either accidentally through exposure or intentionally in the case of potential nanomedical applications. Data from animal experiments cannot be extrapolated to humans because the placenta is the most species-specific mammalian organ 1. Therefore, the ex vivo dual recirculating human placental perfusion, developed by Panigel et al. in 1967 2 and continuously modified by Schneider et al. in 1972 3, can serve as an excellent model to study the transfer of xenobiotics or particles.
Here, we focus on the ex vivo dual recirculating human placental perfusion protocol and its further development to acquire reproducible results.
The placentae were obtained after informed consent of the mothers from uncomplicated term pregnancies undergoing caesarean delivery. The fetal and maternal vessels of an intact cotyledon were cannulated and perfused at least for five hours. As a model particle fluorescently labelled polystyrene particles with sizes of 80 and 500 nm in diameter were added to the maternal circuit. The 80 nm particles were able to cross the placental barrier and provide a perfect example for a substance which is transferred across the placenta to the fetus while the 500 nm particles were retained in the placental tissue or maternal circuit. The ex vivo human placental perfusion model is one of few models providing reliable information about the transport behavior of xenobiotics at an important tissue barrier which delivers predictive and clinical relevant data.

Determination of the Transport Rate of Xenobiotics and Nanomaterials Across the Placenta using the ex vivo Human Placental Perfusion Model

The ex vivo dual recirculating human placental perfusion model can be used to investigate the transfer of xenobiotics and nanoparticles across the human placenta. In this video protocol we describe the equipment and techniques required for a successful execution of a placenta perfusion.

Decades ago the human placenta was thought to be an impenetrable barrier between mother and unborn child. However, the discovery of thalidomide-induced ...

Isolation and Characterization Of Chimeric Human Fc-expressing Proteins Using Protein A Membrane Adsorbers And A Streamlined Workflow

Laboratory scale to industrial scale purification of biomolecules from cell culture supernatants and lysed cell solutions can be accomplished using affinity chromatography.&#160;While affinity chromatography using porous protein A agarose beads packed in columns is arguably the most common method of laboratory scale isolation of antibodies and recombinant proteins expressing Fc fragments of IgG, it can be a time consuming and expensive process.&#160;Time and financial constraints are especially daunting in small basic science labs that must recover hundreds of micrograms to milligram quantities of protein from dilute solutions, yet lack access to high pressure liquid delivery systems and/or personnel with expertise in bioseparations. Moreover, product quantification and characterization may also excessively lengthen processing time over several workdays and inflate expenses (consumables, wages, etc.).&#160;Therefore, a fast, inexpensive, yet effective protocol is needed for laboratory scale isolation and characterization of antibodies and other proteins possessing an Fc fragment.&#160;To this end, we have devised a protocol that can be completed by limited-experience technical staff in less than 9 hr (roughly one workday) and as quickly as 4 hr, as opposed to traditional methods that demand 20+ work hours. Most required equipment is readily available in standard biomedical science, biochemistry, and (bio)chemical engineering labs, and all reagents are commercially available.&#160;To demonstrate this protocol, representative results are presented in which chimeric&#160;murine galectin-1 fused to human Fc (Gal-1hFc) from cell culture supernatant was isolated using a protein A membrane adsorber.&#160;Purified Gal-1hFc was quantified using an expedited Western blotting analysis procedure and characterized using flow cytometry. The streamlined workflow can be modified for other Fc-expressing proteins, such as antibodies, and/or altered to incorporate alternative quantification and characterization methods.

Compared with traditional affinity chromatography using protein A agarose bead-packed columns, protein A membrane adsorbers can significantly speed laboratory-scale isolation of antibodies and other Fc fragment-expressing proteins.&#160;Appropriate analysis and quantification methods can further accelerate protein processing, allowing isolation/characterization to be completed in one workday, instead of 20+ work hours.

Laboratory scale to industrial scale purification of biomolecules from cell culture supernatants and lysed cell solutions can be accomplished using ...

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 ...

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Sandwich-like Microenvironments to Harness Cell/Material Interactions

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. However in vivo, most of the cells are completely surrounded by the extracellular matrix (ECM), resulting in a three-dimensional (3D) distribution of receptors. This may trigger differences in the outside-in signaling pathways and thus in cell behavior.
This article shows that stimulating the dorsal receptors of cells already adhered to a 2D substrate by overlaying a film of a new material (a sandwich-like culture) triggers important changes with respect to standard 2D cultures. Furthermore, the simultaneous excitation of ventral and dorsal receptors shifts cell behavior closer to that found in 3D environments. Additionally, due to the nature of the system, a sandwich-like culture is a versatile tool that allows the study of different parameters in cell/material interactions, e.g., topography, stiffness and different protein coatings at both the ventral and dorsal sides. Finally, since sandwich-like cultures are based on 2D substrates, several analysis procedures already developed for standard 2D cultures can be used normally, overcoming more complex procedures needed for 3D systems.

The following protocol describes the procedure to assemble sandwich-like cultures to be used as an intermediate stage between bi-dimensional (2D) and three-dimensional (3D) cellular environments. The engineered systems can have applications in microscopy, biomechanics, biochemistry and cell biology assays.

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

DNA Nanotubes as a Versatile Tool to Study Semiflexible Polymers

Mechanical properties of complex, polymer-based soft matter, such as cells or biopolymer networks, can be understood in neither the classical frame of flexible polymers nor of rigid rods. Underlying filaments remain outstretched due to their non-vanishing backbone stiffness, which is quantified via the persistence length (lp), but they are also subject to strong thermal fluctuations. Their finite bending stiffness leads to unique, non-trivial collective mechanics of bulk networks, enabling the formation of stable scaffolds at low volume fractions while providing large mesh sizes. This underlying principle is prevalent in nature (e.g., in cells or tissues), minimizing the high molecular content and thereby facilitating diffusive or active transport. Due to their biological implications and potential technological applications in biocompatible hydrogels, semiflexible polymers have been subject to considerable study. However, comprehensible investigations remained challenging since they relied on natural polymers, such as actin filaments, which are not freely tunable. Despite these limitations and due to the lack of synthetic, mechanically tunable, and semiflexible polymers, actin filaments were established as the common model system. A major limitation is that the central quantity lp cannot be freely tuned to study its impact on macroscopic bulk structures. This limitation was resolved by employing structurally programmable DNA nanotubes, enabling controlled alteration of the filament stiffness. They are formed through tile-based designs, where a discrete set of partially complementary strands hybridize in a ring structure with a discrete circumference. These rings feature sticky ends, enabling the effective polymerization into filaments several microns in length, and display similar polymerization kinetics as natural biopolymers. Due to their programmable mechanics, these tubes are versatile, novel tools to study the impact of lp on the single-molecule as well as the bulk scale. In contrast to actin filaments, they remain stable over weeks, without notable degeneration, and their handling is comparably straightforward.

Semiflexible polymers display unique mechanical properties that are extensively applied by living systems. However, systematic studies on biopolymers are limited since properties such as polymer rigidity are inaccessible. This manuscript describes how this limitation is circumvented by programmable DNA nanotubes, enabling experimental studies on the impact of filament rigidity.

Mechanical properties of complex, polymer-based soft matter, such as cells or biopolymer networks, can be understood in neither the classical frame of ...