The authors thank Dr. Kent Seeley, Director of The Center for Drug Discovery and Innovation (CDDI) Proteomics Core Facility at University of South Florida for the use of this facility. This work was supported by NIH/NIDCD grant R01 DC004295 to B.H.A.S.

Ethics Statement
Experiments using mice tissue were approved by the University of South Florida Institutional Animal Care and Use Committee (Protocols 3931R, 3482R) as set forth under the guidelines of the National Institutes of Health.
1. Protein Extraction
<ol>
	<li>Isolate cochlear sensory epithelium from 16 30-day-old (P30) CBA/J mice and store at -80 &#176;C.</li>
	<li>On the day of the experiment, wash tissue with 500 &#181;l of 1x phosphate buffered saline (PBS). Centrifuge for 3 min at 1,000 x g, and remove the supernatant. Repeat for a total of three washes.</li>
	<li>Sonicate tissue for 30 sec on ice in 100 &#956;l of lysis buffer containing 50 mM Tris-HCl, pH 8.0, 120 mM NaCl, 50 mM NaF, 5 mM EDTA, 500 &#956;g/ml AEBSF, 10 &#956;g/ml leupeptin, 100 &#956;g/ml pepstatin, 2 &#956;g/ml aprotinin, and 0.5 &#956;g/ml microcystin using a sonic dismembrator (Model 100; Thermo Fisher). Cool lysate on ice for 1 min between each sonication. Sonicate a total of 3x.</li>
	<li>Centrifuge the extract at 750 x g at 4 &#176;C for 2 min and remove the supernatant to a new microtube. Extract the pellet in 50 &#956;l of lysis buffer by sonicating for 30 sec on ice. Centrifuge the extract at 750 x g at 4 &#176;C for 2 min. Combine both lysates and centrifuge at 28,600 x g at 4 &#176;C for 60 min. Remove the supernatant to a new microtube and add 20 &#956;l lysis buffer containing 0.1% ASB-14 to the pellet. Vortex for 1 min and incubate for 60 min at 4 &#176;C.</li>
	<li>Heat the sample at 95 &#176;C for 5 min, then cool at 4 &#176;C for 60 min. Follow with centrifugation at 16,000 x g at 25 &#176;C for 10 min. Collect the supernatant and transfer to a new tube.</li>
	<li>Centrifuge the suspension at 16,000 x g at 4 &#176;C for 5 min and retain the supernatant for digestion.</li>
</ol>
2. Double Tryptic Protein Digestion of Whole Lysate Using FASP
<ol>
	<li>Add a 30 &#956;l aliquot (&#8804;400 &#956;g) of cochlear protein extract, containing 4% sodium dodecyl sulfate (SDS), 100 mM Tris-HCl, pH 7.6 and 0.1 M dithiothreitol (DTT) directly to a 30K spin filter and mix with 200 &#956;l of 8 M urea in Tris-HCl. Centrifuge at 14,000 x g for 15 min.</li>
	<li>Dilute the concentrate with 200 &#956;l of 8 M urea solution and centrifuge at 14,000 x g for 15 min.</li>
	<li>Add 10 &#181;l of 10x iodoacetamide (IAA) in 8 M urea solution to the concentrate in the filter and vortex for 1 min. Incubate the spin filter for 20 min at room temperature (RT) in the dark followed by centrifugation at 14,000 x g for 10 min.</li>
	<li>Add 100 &#181;l of 8 M urea solution to the concentrate on the filter unit and centrifuge at 14,000 x g for 15 min. Repeat this step 2x. Add 100 &#956;l of 50 mM ammonium bicarbonate (ABC) solution to the filter unit and centrifuge at 14,000 x g for 10 min. Repeat this step 2x.</li>
	<li>Add 0.4 &#956;g/&#956;l of trypsin in 1:100 (w/w) enzyme-to-protein ratio and incubate overnight (O/N) at 37 &#176;C.</li>
	<li>Following incubation, add 40 &#956;l of 50 mM ABC solution to filter unit and centrifuge at 14,000 x g for 10 min. Repeat this step 1x.</li>
	<li>Add 50 &#956;l of 0.5 M NaCl solution to the spin filter and centrifuge at 14,000 x g for 10 min. Transfer the filtrate containing the tryptic peptides to a fresh microtube and acidify with formic acid (FA) to 1.0% to stop the digestion.</li>
	<li>Wash the filter unit with 40 &#181;l of 8 M urea, and then wash 2x with 40 &#181;l 18 M&#937; water.</li>
	<li>Wash the filter unit 3x with 100 &#956;l of 50 mM ABC solution. After the final wash add 0.4 &#956;g/&#956;l of trypsin in 1:100 (w/w) enzyme-to-protein ratio and incubate O/N at 37 &#176;C.</li>
	<li>Elute tryptic peptides from the second digest by adding 40 &#956;l of 50 mM ABC solution to the filter unit and centrifuge at 14,000 x g for 10 min. Repeat this step 1x.</li>
	<li>Add 50 &#956;l of 0.5 M NaCl solution to the filter unit and centrifuge at 14,000 x g for 10 min. Transfer the filtrate containing the tryptic peptides to a fresh microtube and acidify with FA to 1.0%.</li>
	<li>The samples can be quantified using the microplate colorimetric assay using BSA as the standard.</li>
</ol>
3. Endoproteinase LysC and Tryptic Protein Digestion of Whole Lysate Using FASP
<ol>
	<li>Add a 30 &#956;l aliquot (&#8804;400 &#956;g) of cochlear protein extract containing 4% SDS, 100 mM Tris-HCl, pH 7.6 and 0.1 M DTT directly to a 30K spin filter and mix with 200 &#956;l of 8 M urea in Tris-HCl. Centrifuge at 14,000 x g for 15 min.</li>
	<li>Dilute the concentrate with 200 &#956;l of 8 M urea solution and centrifuge at 14,000 x g for 15 min.</li>
	<li>Add 10 &#956;l of 10x IAA in 8 M urea solution to the concentrate in the filter and vortex for 1 min. Incubate the spin filter for 20 min at RT in the dark then centrifuge at 14,000 x g for 10 min.</li>
	<li>Add 100 &#956;l of 8 M urea solution to the concentrate on the filter unit and centrifuge at 14,000 x g for 15 min. Repeat this step 2x. Add 100 &#956;l of 100 mM ABC solution to the filter unit and centrifuge at 14,000 x g for 10 min. Repeat this step 2x.</li>
	<li>Add 0.1 &#956;g/&#956;l of endoproteinase LysC in a 1:50 (w/w) enzyme-to-protein ratio and incubate O/N at 30 &#176;C.</li>
	<li>Following incubation, add 40 &#956;l of 100 mM ABC solution to the filter unit and centrifuge at 14,000 x g for 10 min. Repeat this step 1x.</li>
	<li>Add 50 &#956;l of 0.5 M NaCl solution to the spin filter and centrifuge at 14,000 x g for 10 min. Transfer the filtrate containing the LysC peptides to a fresh microtube and acidify with FA to 1.0% to stop the digestion.</li>
	<li>Wash the filter unit with 40 &#956;l of 8 M urea, and then wash 2x with 40 &#956;l 18 M&#937; water.</li>
	<li>Wash the filter unit 3x with 100 &#956;l of 50 mM ABC solution. After the final wash add 0.4 &#956;g/&#956;l of trypsin in 1:100 enzyme-to-protein ratio and incubate O/N at 37 &#176;C.</li>
	<li>Elute tryptic peptides by adding 40 &#956;l of 50 mM ABC solution to the filter unit and centrifuge at 14,000 x g for 10 min. Repeat this step 1x.</li>
	<li>Add 50 &#181;l of 0.5 M NaCl solution to the filter unit and centrifuge at 14,000 x g for 10 min. Transfer the filtrate containing the tryptic peptides to a fresh microtube and acidify with 1.0% FA.</li>
	<li>The sample can be quantified using the microplate colorimetric assay with BSA as the standard.</li>
</ol>
4. Desalting Peptides Using Spin Columns
<ol>
	<li>Activate a C18 MacroSpin column&#160;by adding 500 &#181;l of acetonitrile (ACN) and centrifuge at 1,100 x g for 1 min. Discard the flow-through after centrifugation.</li>
	<li>Equilibrate the column with 500 &#956;l of 0.1% FA and centrifuge at 1,100 x g for 1 min. Discard the flow-through and repeat this step 1x.</li>
	<li>Add up to 500 &#956;l of the peptide digest to the column and centrifuge at 1,100 x g for 1 min. If the sample volume is greater than 500 &#181;l then repeat this step.</li>
	<li>Wash the column with 500 &#956;l of 0.1% FA and centrifuge at 1,100 x g for 1 min. Discard the flow-through. Repeat this step 1x.</li>
	<li>Add 250 &#181;l of a 90:10 ACN-to-water ratio to the column and centrifuge at 1,100 x g for 1 min. Collect the eluent containing the desalted peptides and transfer to a fresh microtube. Repeat this step 1x.</li>
	<li>Dry the desalted peptide sample in a vacuum centrifuge and avoid letting the sample dry completely.</li>
</ol>
5. Ion Exchange Chromatography
<ol>
	<li>Inject 50-100 &#956;g of digested protein sample onto a 200 x 2.1 mm, 5 &#956;m SCX column (Polysulfoethyl A) to separate peptides.</li>
	<li>Use a gradient of 2-40% B over 50 min with a flow rate of 250 &#956;l/min. Solvent A is 5 mM ammonium formate, pH 3.0 in 25% acetonitrile and 75% ddH2O. Solvent B is 500 mM ammonium formate, pH 6.0 in 25% ACN and 75% ddH2O.</li>
	<li>Monitor the peptide fractions at 280 nm and collect the fractions in 2 min intervals using a fraction collector.</li>
	<li>Dry fractions in a vacuum concentrator and resuspend in 500 &#956;l of 50% acetonitrile in ddH2O containing 5% FA to assist with salt removal.</li>
	<li>Redry fractions and store at -80 &#176;C until ready to use for nano LC-MS/MS analysis.</li>
</ol>
6. Acetone Precipitation
Prior to GELFrEE separation the cochlear protein supernatant has to be desalted. Acetone precipitation can be used to desalt and concentrate proteins.
<ol>
	<li>Add three volumes of ice-cold acetone to the cochlear supernatant. Gently vortex and incubate at -20 &#176;C O/N.</li>
	<li>Centrifuge the sample mixture at 15,000 x g for 15 min at 4 &#176;C and remove the supernatant.</li>
	<li>Wash the protein pellet 3x with chilled acetone and centrifuge at 14,000 x g for 5 min at 4 &#176;C.</li>
	<li>Air-dry the pellet and dissolve in 112 &#956;l of 18 M&#937; water.</li>
	<li>Determine the protein concentration using the microplate colorimetric assay.</li>
</ol>
7. GELFrEE Fractionation of Cochlear Sensory Epithelium
<ol>
	<li>Add 30 &#956;l of 5x sample buffer (0.25 M Tris-HCl pH 6.8, 10% (w/v) SDS, 50% glycerol, 0.5% (w/v) bromophenol blue) to the desalted protein suspended in 112 &#956;l of 18 M&#937; water.</li>
	<li>Add 8 &#956;l of 1M DTT and heat at 95 &#176;C for 5 min to reduce protein disulfide bonds. After heating allow cooling to room temperature.</li>
	<li>Add 8 ml of running buffer to anode reservoir, 150 &#956;l of running buffer to sample collection chamber, and 6 ml of running buffer to cathode reservoir.</li>
	<li>Remove any buffer that may have flowed into the sample-loading chamber using a pipette.</li>
	<li>Load 150 &#956;l (&#8804;1 mg) of the cochlear protein mixture, into the sample-loading chamber using an 8% Tris-acetate cartridge with a mass range of 3.5-150 kDa.</li>
	<li>Lower electrodes and close instrument lid to start the run.</li>
	<li>At the first paused time on the instrument add 2 ml of running buffer to the cathode reservoir and resume run.</li>
	<li>At each paused interval on the instrument, collect the sample from the sample collection chamber using a pipette and add to a freshly labeled microtube. Wash the collection chamber 2x with 150 &#956;l of running buffer and discard.</li>
	<li>Add 150 &#956;l of fresh running buffer to the sample collection chamber and resume run. Collect the protein fractions over a time period of 2.6 hr in a total volume of 150 &#956;l/fraction.</li>
</ol>
8. 1D Gel Electrophoresis of GELFrEE Fractions
1D gel electrophoresis can be used to visualize the results from GELFrEE fractionation prior to enzymatic digestion and MS analysis. GELFrEE protein fractions can be separated on a 4-15% Tris-HCl gel.
<ol>
	<li>Mix a 5 &#956;l aliquot of each GELFrEE fraction with 5 &#181;l of sample diluting buffer (350 mM DTT in sample buffer).</li>
	<li>Heat the samples at 95 &#176;C for 3 min.</li>
	<li>Cool samples to RT.</li>
	<li>Load 10 &#956;l of each GELFrEE fraction in individual gel lanes and load 5 &#956;l of molecular weight standard in the last unused lane.</li>
	<li>Run the gel at 125 V for 1.5 hr or until the dye front reaches the bottom of the gel.</li>
	<li>Carefully remove the gel and stain with Silver Stain Plus to visualize the protein separation.</li>
</ol>
9. Protein Digestion of GELFrEE Fractions Using FASP
A modified FASP procedure is used for detergent removal and digestion of the GELFrEE fractions.
<ol>
	<li>Add each individual fraction directly to a 30K filter unit; mix with 200 &#956;l of 8 M urea in Tris-HCl and centrifuge at 14,000 x g for 25 min.</li>
	<li>Dilute the concentrate with 200 &#956;l of urea solution and centrifuge at 14,000 x g for 12 min. Repeat this step 1x.</li>
	<li>Add 10 &#956;l of 10x IAA in urea solution to the concentrate in each filter unit, vortex for 1 min, and incubate for 30 min at RT in the dark.</li>
	<li>Add 100 &#956;l of urea solution to the concentrate on the filter units and centrifuge at 14,000 x g for 15 min. Repeat this step 2x. Add 100 &#956;l of 100 mM ABC solution to the filter units and centrifuge at 14,000 x g for 10 min. Repeat this step 2x.</li>
	<li>Add 0.1 &#956;g/&#956;l of LysC for a 1:50 (w/w) enzyme-to-protein ratio to the filter units and incubate O/N at 30 &#176;C.</li>
	<li>Following incubation, add 40 &#956;l of 100 mM ABC solution to the spin filters and centrifuge at 14,000 x g for 10 min. Repeat this step 1x.</li>
	<li>Add 50 &#956;l of 0.5 M NaCl solution to the spin filters and centrifuge at 14,000 x g for 10 min. Transfer the filtrate containing the LysC peptides from each spin filter to a fresh microtube and acidify with trifluoroacetic acid (TFA).</li>
	<li>Wash the spin filters with 40 &#956;l of 8 M urea, then wash 2x with 40 &#956;l 18 M&#937; water.</li>
	<li>Wash the spin filters 3x with 100 &#956;l of 50 mM ABC solution. After the final wash add 0.4 &#956;g/&#956;l of trypsin in a 1:100 (w/w) enzyme-to-protein ratio and incubate O/N at 37 &#176;C.</li>
	<li>Elute tryptic peptides from each filter unit by adding 40 &#956;l of 50 mM ABC solution and centrifuge at 14,000 x g for 10 min. Repeat this step 1x.</li>
	<li>Add 50 &#956;l of 0.5 M NaCl solution to the filter units and centrifuge at 14,000 x g for 10 min. Transfer the filtrate containing the tryptic peptides from each filter unit to a fresh microtube and acidify with TFA.</li>
	<li>Dry all digests in a vacuum concentrator.</li>
</ol>
10. Sample Preparation for LC-MS/MS
<ol>
	<li>Reconstitute dried LysC and tryptic peptide digests fractions in 20 &#181;l of 0.1% FA and vortex.</li>
	<li>Centrifuge samples at 20,000 x g for 10 min and remove the top 95% of sample to a new sample vial.</li>
	<li>Inject 5 &#181;l of each peptide fraction onto a 100 &#956;m x 25 mm sample trap to remove salts and contaminants and perform chromatographic separation on a 75 &#956;m &#215; 10 cm C18 column.</li>
	<li>Use a gradient of 2-40% B over 100 min with a flow rate of 200 nl/min. Solvent A is 95% ddH2O and 5% acetonitrile containing 0.1% FA. Solvent B is 80% ACN and 20% ddH2O containing 0.1% FA.</li>
	<li>Collect ten tandem mass spectra for each MS scan on a high-resolution mass spectrometer. An LTQ Orbitrap mass spectrometer was used in this experiment.</li>
</ol>
11. Protein Identification
<ol>
	<li>Identify proteins using the UniProt mouse database containing both forward and reverse protein sequences and common contaminants. MaxQuant&#160;software with MASCOT search engine is used to search the protein database.</li>
	<li>The search parameters that can be used include; fixed modification of carbamidomethyl of cysteine, variable modifications of oxidation of methionine, protein N-terminal acetylation, maximum of 2 missed cleavage. The minimum peptide length to consider for identification is 6 amino acids. Enter a peptide mass concentration error of &#177;8 ppm and a fragment mass tolerance of 1.2 Da (mass error is dependent on the mass spectrometer being used).</li>
	<li>In the MaxQuant software, use a 1% false discovery rate (FDR) for peptide and protein identification. Protein identification is listed with the number of matched peptides, the percent protein sequence coverage, and the peptide sequences.</li>
</ol>

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R., Zougman, A., Nagaraj, N., Mann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Universal+sample+preparation+method+for+proteome+analysis.">Universal sample preparation method for proteome analysis.</a> Nat. Meth. 6, 359-362 (2009).</li><li>Wisniewski, J. R., Zielinska, D. F., Mann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+ultrafiltration+units+for+proteomic+and+N-glycoproteomic+analysis+by+the+filter-aided+sample+preparation+method.">Comparison of ultrafiltration units for proteomic and N-glycoproteomic analysis by the filter-aided sample preparation method.</a> Anal. Biochem. 410, 307-309 (2011).</li><li>Darville, L. N., Sokolowski, B. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In-depth+Proteomic+Analysis+of+Mouse+Cochlear+Sensory+Epithelium+by+Mass+Spectrometry.">In-depth Proteomic Analysis of Mouse Cochlear Sensory Epithelium by Mass Spectrometry.</a> J Proteome Res. 12 (8), 3620-3630 (2013).</li><li>Vizcaino, J. A., Cote, R. G., Csordas, A., Dianes, J. A., Fabregat, A., Foster, J. M., Griss, J., Alpi, E., Birim, M., Contell, J., O'Kelly, G., Schoenegger, A., Ovelleiro, D., Perez-Riverol, Y., Reisinger, F., Rios, D., Wang, R., Hermjakob, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+PRoteomics+IDEntifications+(PRIDE)+database+and+associated+tools:+status+in+2013.">The PRoteomics IDEntifications (PRIDE) database and associated tools: status in 2013.</a> Nucleic Acids Res. 41, 1063-1069 (2013).</li></ol>

The authors declare no competing interests.

The key steps to maximizing protein identification from the cochlear sensory epithelium are: 1) use of multiple endoproteinases for digestion, 2) use of multiple separation techniques, and 3) utilization of a high-resolution mass spectrometer. The application of multiple enzymes increases the number of peptides and improves protein sequence coverage, hence improving the number of identified proteins from the cochlear tissue. Trypsin, the most commonly used protease provides efficient and specific cleavage of proteins, ge...

Proteomics is the study of complex biological systems by analyzing protein expression, function, modifications, and interactions1. Several methods have been utilized for proteome analysis of the inner ear, including antibody microarray2, two-dimensional gel electrophoresis3-5, and DIGE6. However, only a limited number of proteins have been identified and characterized2,7-10, compared to the over 10,000 genes and expressed sequence tags (ESTs) identified in the inner ear11,12, MS is the most commonly used and comprehensive technique in proteomics for protein characterization. Analysis of complex proteomic samples, such as the cochlea, can be challenging. However, the combination of multiple separation techniques with MS enables the identification of a greater number of peptides and proteins, due to an increased dynamic concentration range and peak capacity13. Multidimensional chromatography reduces highly complex protein mixtures by allowing the use of different adsorption mechanisms. There are two commonly used MS proteome analysis approaches, shotgun and bottom-up proteomics. In shotgun proteomics, a mixture of intact proteins is enzymatically digested and separated using multidimensional chromatography with strong cation-exchange chromatography (SCX) followed by reversed-phase liquid chromatography (RPLC)14,15. The separated peptides are subjected to tandem MS and database searching15. A major advantage of this technique is that thousands of proteins can be identified in a single analysis and the technique is better suited to membrane proteins.
In the bottom-up approach, the protein mixture is separated, usually by one- or two-dimensional electrophoresis, and the individual protein bands or spots cut out and digested with an enzyme such as trypsin, usually resulting in multiple peptides. However, another more recently developed electrophoretic approach, used in bottom-up proteomics, is GELFrEE. This technique fractionates protein samples in liquid-phase and makes them less complex prior to analysis. This technique is reproducible, offers high protein recovery, and reduces the distribution of high abundant proteins in complex protein samples16. Peptides, resulting from separated proteins, are analyzed by MS, by using peptide mass fingerprinting or tandem MS (MS/MS), to create sequence tags for database searching17-19. Some of the major advantages of using the bottom-up approach are the ability to obtain high-resolution separations and comprehensive protein coverage. Bottom-up proteomics is the most widely used technique in proteomics20, hence, several bioinformatics tools are available. In addition, proteins can be separated in a complex mixture before digestion, so there is a greater chance of identification.
One of the major challenges in using the inner ear for proteomic analysis is its small size, restricted accessibility, and cell type diversity21. In addition, key proteins that distinguish its functionality, such as ion channels, transporters and receptors, are membrane proteins, which can be difficult to isolate22. Thus, filter-aided sample preparation (FASP) is advantageous for proteomic analyses of tissues that are limited for protein extraction and that require detergents to solubilize membranes23. This filtering allows for the MS analysis of membrane and soluble proteins and for the ability to isolate peptides from low molecular weight contaminants23,24.
The present protocol describes commonly used proteomic approaches that are combined and modified to analyze both soluble and membrane proteins and to maximize the number of protein IDs from the cochlear sensory epithelium. We will describe using shotgun proteomics with FASP multi-digestion, ion exchange chromatography, high resolution MS, and data analysis. In addition, we will describe bottom-up proteomics with GELFrEE, FASP multi-digestion, high resolution MS, and data analysis.

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			<td height="20" style="height:20px;width:264px;">8% Tris-acetate cartridge&#160;</td>
			<td style="width:159px;">Protein Discovery</td>
			<td style="width:133px;">42103</td>
			<td style="width:193px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Acetone</td>
			<td>Sigma-Aldrich</td>
			<td>179124</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Acetonitrile</td>
			<td>Honeywell</td>
			<td>015-1L</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">AEBSF</td>
			<td>Calbiochem</td>
			<td>101500</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Ammonium formate</td>
			<td>Fisher Scientific</td>
			<td>AC16861</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Aprotinin</td>
			<td>Calbiochem</td>
			<td>616370</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">ASB-14&#160;</td>
			<td>Calbiochem</td>
			<td>182750-5GM</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Bovine serum albumin</td>
			<td>BioRad</td>
			<td>500-0112</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;">C18 column&#160;</td>
			<td>New Objective</td>
			<td>A25112</td>
			<td>75 &#956;m &#215; 10 cm&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DC Protein Assay</td>
			<td>BioRad</td>
			<td>500-0116</td>
			<td>Microplate Assay Protocol</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">EDTA</td>
			<td>Sigma-Aldrich</td>
			<td>E9884</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Endoproteinase Lys-C</td>
			<td>Sigma-Aldrich</td>
			<td>P3428</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">FASP Protein Digestion Kit</td>
			<td>Protein Discovery</td>
			<td>44250</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Formic acid&#160;</td>
			<td>Fluka</td>
			<td>94318</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">GELFrEE Fractionation System</td>
			<td>Protein Discovery</td>
			<td>42001</td>
			<td>GELFrEE 8100</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Leupeptin</td>
			<td>Calbiochem</td>
			<td>108975</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">MacroSpin Column</td>
			<td>The Nest Group</td>
			<td>SMM SS18V</td>
			<td>Silica C18</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Microcystin</td>
			<td>Calbiochem</td>
			<td>475815</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pepstatin</td>
			<td>Sigma-Aldrich</td>
			<td>P5318</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Polysulfoethyl A Column</td>
			<td>The Nest Group</td>
			<td>202SE0503</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium dodecyl sulfate (SDS)</td>
			<td>Sigma-Aldrich</td>
			<td>L3771</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sonic Dismembrator&#160;</td>
			<td>Thermo Fisher</td>
			<td>15-338-53</td>
			<td>Model 100</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Trypsin</td>
			<td>Sigma-Aldrich</td>
			<td>T6567</td>
			<td>Proteomics Grade</td>
		</tr>
	</tbody>
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bottom-up and shotgun proteomics to identify a comprehensive cochlear proteome

Proteomics is a commonly used approach that can provide insights into complex biological systems. The cochlear sensory epithelium contains receptors that transduce the mechanical energy of sound into an electro-chemical energy processed by the peripheral and central nervous systems. Several proteomic techniques have been developed to study the cochlear inner ear, such as two-dimensional difference gel electrophoresis (2D-DIGE), antibody microarray, and mass spectrometry (MS). MS is the most comprehensive and versatile tool in proteomics and in conjunction with separation methods can provide an in-depth proteome of biological samples. Separation methods combined with MS has the ability to enrich protein samples, detect low molecular weight and hydrophobic proteins, and identify low abundant proteins by reducing the proteome dynamic range. Different digestion strategies can be applied to whole lysate or to fractionated protein lysate to enhance peptide and protein sequence coverage. Utilization of different separation techniques, including strong cation exchange (SCX), reversed-phase (RP), and gel-eluted liquid fraction entrapment electrophoresis (GELFrEE) can be applied to reduce sample complexity prior to MS analysis for protein identification.

To obtain the most comprehensive proteome of the cochlear sensory epithelium, quick tissue dissection is required prior to protein extraction and sample preparation. Two proteomic techniques can be used, shotgun and bottom-up proteomics. To prepare samples for shotgun proteomics, FASP digestion procedure was used as illustrated in Figure 1. The FASP method allows for concentration of proteins, removal of detergents, and digestion of proteins using multiple enzymes. There were two double digestion procedu...

Watch this Scientific Journal Video about Bottom-up and Shotgun Proteomics to Identify a Comprehensive Cochlear Proteome at JoVE.com

Bottom-up and Shotgun Proteomics to Identify a Comprehensive Cochlear Proteome

Proteome analysis of the cochlear sensory epithelium can be challenging due to its small size and because membrane proteins are difficult to isolate and identify. Both membrane and soluble proteins can be identified by combining multiple preparative methods and separation techniques along with high-resolution mass spectrometry.

Proteomics is a commonly used approach that can provide insights into complex biological systems. The cochlear sensory epithelium contains receptors that ...

bottom-up-shotgun-proteomics-to-identify-comprehensive-cochlear

Research

JoVE Journal

Biology

18.6K vistas.  University of South Florida.  Análisis del proteoma del epitelio sensorial coclear puede ser un reto debido a su pequeño tamaño y porque las proteínas de membrana son difíciles de aislar e identificar. Tanto la membrana y proteínas solubles se pueden identificar mediante la combinación de múltiples métodos de preparación y técnicas de separación junto con espectrometría de masas de alta resolución. 

Video: Bottom-up and Shotgun Proteomics to Identify a Comprehensive Cochlear Proteome

Isolation and Analysis of Hematopoietic Stem Cells from the Placenta

Hematopoietic stem cells (HSCs) have the ability to self-renew and generate all cell types of the blood lineages throughout the lifetime of an individual. All HSCs emerge during embryonic development, after which their pool size is maintained by self-renewing cell divisions. Identifying the anatomical origin of HSCs and the critical developmental events regulating the process of HSC development has been complicated as many anatomical sites participate during fetal hematopoiesis. Recently, we identified the placenta as a major hematopoietic organ where HSCs are generated and expanded in unique microenvironmental niches (Gekas, et al 2005, Rhodes, et al 2008). Consequently, the placenta is an important source of HSCs during their emergence and initial expansion.
In this article, we show dissection techniques for the isolation of murine placenta from E10.5 and E12.5 embryos, corresponding to the developmental stages of initiation of HSCs and the peak in the size of the HSC pool in the placenta, respectively. In addition, we present an optimized protocol for enzymatic and mechanical dissociation of placental tissue into single-cell suspension for use in flow cytometry or functional assays. We have found that use of collagenase for single-cell suspension of placenta gives sufficient yields of HSCs. An important factor affecting HSC yield from the placenta is the degree of mechanical dissociation prior to, and duration of, enzymatic treatment.
We also provide a protocol for the preparation of fixed-frozen placental tissue sections for the visualization of developing HSCs by immunohistochemistry in their precise cellular niches. As hematopoietic specific antigens are not preserved during preparation of paraffin embedded sections, we routinely use fixed frozen sections for localizing placental HSCs and progenitors.

We have identified the placenta as a major hematopoietic organ during development. We found that hematopoietic stem cells (HSCs) are both generated and expanded in the placenta in unique microenvironmental niches. Here, we describe experimental techniques required for isolation and visualization of HSCs in the mouse placenta.

Hematopoietic stem cells (HSCs) have the ability to self-renew and generate all cell types of the blood lineages throughout the lifetime of an individual. ...

Proteomics to Identify Proteins Interacting with P2X2 Ligand-Gated Cation Channels

Ligand-gated ion channels underlie synaptic communication in the nervous system1. In mammals there are three families of ligand-gated channels: the cys loop, the glutamate-gated and the P2X receptor channels2. In each case binding of transmitter leads to the opening of a pore through which ions flow down their electrochemical gradients. Many ligand-gated channels are also permeable to calcium ions3, 4, which have downstream signaling roles5 (e.g. gene regulation) that may exceed the duration of channel opening. Thus ligand-gated channels can signal over broad time scales ranging from a few milliseconds to days. Given these important roles it is necessary to understand how ligand-gated ion channels themselves are regulated by proteins, and how these proteins may tune signaling. Recent studies suggest that many, if not all, channels may be part of protein signaling complexes6. In this article we explain how to identify the proteins that bind to the C-terminal aspects of the P2X2 receptor cytosolic domain.
P2X receptors are ATP-gated cation channels and consist of seven subunits (P2X1-P2X7). P2X receptors are widely expressed in the brain, where they mediate excitatory synaptic transmission and presynaptic facilitation of neurotransmitter release7. P2X receptors are found in excitable and non-excitable cells and mediate key roles in neuronal signaling, inflammation and cardiovascular function8. P2X2 receptors are abundant in the nervous system9 and are the focus of this study. Each P2X subunit is thought to possess two membrane spanning segments (TM1 &amp; TM2) separated by an extracellular region7 and intracellular N and C termini (Fig 1a)7. P2X subunits10 (P2X1-P2X7) show 30-50% sequence homology at the amino acid level11. P2X receptors contain only three subunits, which is the simplest stoichiometry among ionotropic receptors. The P2X2 C-terminus consists of 120 amino acids (Fig 1b) and contains several protein docking consensus sites, supporting the hypothesis that P2X2 receptor may be part of signaling complexes. However, although several functions have been attributed to the C-terminus of P2X2 receptors9 no study has described the molecular partners that couple to the intracellular side of this protein via the full length C-terminus. In this methods paper we describe a proteomic approach to identify the proteins which interact with the full length C-terminus of P2X2 receptors.

We describe a simple protocol to identify brain proteins that bind to the full length C terminus of ATP-gated P2X2 receptors. The extension and systematic application of this approach to all P2X receptors is expected to lead to a better understanding of P2X receptor signaling.

Ligand-gated ion channels underlie synaptic communication in the nervous system1. In mammals there are three families of ligand-gated channels: the cys ...

PAR-CliP - A Method to Identify Transcriptome-wide the Binding Sites of RNA Binding Proteins

RNA transcripts are subjected to post-transcriptional gene regulation by interacting with hundreds of RNA-binding proteins (RBPs) and microRNA-containing ribonucleoprotein complexes (miRNPs) that are often expressed in a cell-type dependently. To understand how the interplay of these RNA-binding factors affects the regulation of individual transcripts, high resolution maps of in vivo protein-RNA interactions are necessary1.
A combination of genetic, biochemical and computational approaches are typically applied to identify RNA-RBP or RNA-RNP interactions. Microarray profiling of RNAs associated with immunopurified RBPs (RIP-Chip)2 defines targets at a transcriptome level, but its application is limited to the characterization of kinetically stable interactions and only in rare cases3,4 allows to identify the RBP recognition element (RRE) within the long target RNA. More direct RBP target site information is obtained by combining in vivo UV crosslinking5,6 with immunoprecipitation7-9 followed by the isolation of crosslinked RNA segments and cDNA sequencing (CLIP)10. CLIP was used to identify targets of a number of RBPs11-17. However, CLIP is limited by the low efficiency of UV 254 nm RNA-protein crosslinking, and the location of the crosslink is not readily identifiable within the sequenced crosslinked fragments, making it difficult to separate UV-crosslinked target RNA segments from background non-crosslinked RNA fragments also present in the sample.
We developed a powerful cell-based crosslinking approach to determine at high resolution and transcriptome-wide the binding sites of cellular RBPs and miRNPs that we term PAR-CliP (Photoactivatable-Ribonucleoside-Enhanced Crosslinking and Immunoprecipitation) (see Fig. 1A for an outline of the method). The method relies on the incorporation of photoreactive ribonucleoside analogs, such as 4-thiouridine (4-SU) and 6-thioguanosine (6-SG) into nascent RNA transcripts by living cells. Irradiation of the cells by UV light of 365 nm induces efficient crosslinking of photoreactive nucleoside-labeled cellular RNAs to interacting RBPs. Immunoprecipitation of the RBP of interest is followed by isolation of the crosslinked and coimmunoprecipitated RNA. The isolated RNA is converted into a cDNA library and deep sequenced using Solexa technology. One characteristic feature of cDNA libraries prepared by PAR-CliP is that the precise position of crosslinking can be identified by mutations residing in the sequenced cDNA. When using 4-SU, crosslinked sequences thymidine to cytidine transition, whereas using 6-SG results in guanosine to adenosine mutations. The presence of the mutations in crosslinked sequences makes it possible to separate them from the background of sequences derived from abundant cellular RNAs.
Application of the method to a number of diverse RNA binding proteins was reported in Hafner et al.18

RNA transcripts are subject to extensive posttranscriptional regulation that is mediated by a multitude of trans-acting RNA-binding proteins (RBPs). Here we present a generalizable method to identify precisely and on a transcriptome-wide scale the RNA binding sites of RBPs.

RNA transcripts are subjected to post-transcriptional gene regulation by interacting with hundreds of RNA-binding proteins (RBPs) and microRNA-containing ...

Quantification of Proteins Using Peptide Immunoaffinity Enrichment Coupled with Mass Spectrometry

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in quantitation, are associated with a high cost, long lead time, and are fraught with drawbacks (e.g. heterophilic antibodies, autoantibody interference, 'hook-effect').1 An alternative technique is affinity enrichment of peptides coupled with quantitative mass spectrometry, commonly referred to as SISCAPA (Stable Isotope Standards and Capture by Anti-Peptide Antibodies).2 In this technique, affinity enrichment of peptides with stable isotope dilution and detection by selected/multiple reaction monitoring mass spectrometry (SRM/MRM-MS) provides quantitative measurement of peptides as surrogates for their respective proteins. SRM/MRM-MS is well established for accurate quantitation of small molecules 3, 4 and more recently has been adapted to measure the concentrations of proteins in plasma and cell lysates.5-7 To achieve quantitation of proteins, these larger molecules are digested to component peptides using an enzyme such as trypsin. One or more selected peptides whose sequence is unique to the target protein in that species (i.e. "proteotypic" peptides) are then enriched from the sample using anti-peptide antibodies and measured as quantitative stoichiometric surrogates for protein concentration in the sample. Hence, coupled to stable isotope dilution (SID) methods (i.e. a spiked-in stable isotope labeled peptide standard), SRM/MRM can be used to measure concentrations of proteotypic peptides as surrogates for quantification of proteins in complex biological matrices. The assays have several advantages compared to traditional immunoassays. The reagents are relatively less expensive to generate, the specificity for the analyte is excellent, the assays can be highly multiplexed, enrichment can be performed from neat plasma (no depletion required), and the technique is amenable to a wide array of proteins or modifications of interest.8-13 In this video we demonstrate the basic protocol as adapted to a magnetic bead platform.

Stable Isotope Standards and Capture by Anti-Peptide Antibodies (SISCAPA) couples affinity enrichment of peptides with stable isotope dilution mass spectrometry (MRM-MS) to provide quantitative measurement of peptides as surrogates for their respective proteins. Here we describe the protocol using magnetic particles in a partially automated format.

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in ...

Orthogonal Protein Purification Facilitated by a Small Bispecific Affinity Tag

Due to the high costs associated with purification of recombinant proteins the protocols need to be rationalized. For high-throughput efforts there is a demand for general methods that do not require target protein specific optimization1 . To achieve this, purification tags that genetically can be fused to the gene of interest are commonly used2 . The most widely used affinity handle is the hexa-histidine tag, which is suitable for purification under both native and denaturing conditions3 . The metabolic burden for producing the tag is low, but it does not provide as high specificity as competing affinity chromatography based strategies1,2.
Here, a bispecific purification tag with two different binding sites on a 46 amino acid, small protein domain has been developed. The albumin-binding domain is derived from Streptococcal protein G and has a strong inherent affinity to human serum albumin (HSA). Eleven surface-exposed amino acids, not involved in albumin-binding4 , were genetically randomized to produce a combinatorial library. The protein library with the novel randomly arranged binding surface (Figure 1) was expressed on phage particles to facilitate selection of binders by phage display technology. Through several rounds of biopanning against a dimeric Z-domain derived from Staphylococcal protein A5, a small, bispecific molecule with affinity for both HSA and the novel target was identified6 .
The novel protein domain, referred to as ABDz1, was evaluated as a purification tag for a selection of target proteins with different molecular weight, solubility and isoelectric point. Three target proteins were expressed in Escherishia coli with the novel tag fused to their N-termini and thereafter affinity purified. Initial purification on either a column with immobilized HSA or Z-domain resulted in relatively pure products. Two-step affinity purification with the bispecific tag resulted in substantial improvement of protein purity. Chromatographic media with the Z-domain immobilized, for example MabSelect SuRe, are readily available for purification of antibodies and HSA can easily be chemically coupled to media to provide the second matrix.
This method is especially advantageous when there is a high demand on purity of the recovered target protein. The bifunctionality of the tag allows two different chromatographic steps to be used while the metabolic burden on the expression host is limited due to the small size of the tag. It provides a competitive alternative to so called combinatorial tagging where multiple tags are used in combination1,7.

A novel and highly efficient two-step affinity chromatography protocol has been developed and is described in detail. The method is based on a small purification tag with two inherent affinities and is applicable to a wide range of target proteins with different properties.

Due to the high costs associated with purification of recombinant proteins the protocols need to be rationalized. For high-throughput efforts there is a ...

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated tools and resources for studies in this system are facilitating continued discovery of genes with more subtle phenotypes or roles. 
Here we present a generalized protocol we adapted for identifying C. elegans genes with postembryonic phenotypes of interest using RNAi2. This procedure is easily modified to assay the phenotype of choice, whether by light or fluorescence optics on a dissecting or compound microscope. This screening protocol capitalizes on the physical assets of the organism and molecular tools the C. elegans research community has produced. As an example, we demonstrate the use of an integrated transgene that expresses a fluorescent product in an RNAi screen to identify genes required for the normal localization of this product in late stage larvae and adults. First, we used a commercially available genomic RNAi library with full-length cDNA inserts. This library facilitates the rapid identification of multiple candidates by RNAi reduction of the candidate gene product. Second, we generated an integrated transgene that expresses our fluorecently tagged protein of interest in an RNAi-sensitive background. Third, by exposing hatched animals to RNAi, this screen permits identification of gene products that have a vital embryonic role that would otherwise mask a post-embryonic role in regulating the protein of interest. Lastly, this screen uses a compound microscope equipped for single cell resolution.

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

We describe a sensitized method to identify postembryonic regulators of protein expression and localization in C. elegans using an RNAi-based genomic screen and an integrated transgene that expresses a functional, fluorescently tagged protein.

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated ...

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the messagefor review see 1. However, it's now widely accepted that synonymous codon usage is the primary cause of non-uniform translational elongation rates1. Synonymous codons are not used with identical frequency. A bias exists in the use of synonymous codons with some codons used more frequently than others2. Codon bias is organism as well as tissue specific2,3. Moreover, frequency of codon usage is directly proportional to the concentrations of cognate tRNAs4. Thus, a frequently used codon will have higher multitude of corresponding tRNAs, which further implies that a frequent codon will be translated faster than an infrequent one. Thus, regions on mRNA enriched in rare codons (potential pause sites) will as a rule slow down ribosome movement on the message and cause accumulation of nascent peptides of the respective sizes5-8. These pause sites can have functional impact on the protein expression, mRNA stability and protein foldingfor review see 9. Indeed, it was shown that alleviation of such pause sites can alter ribosome movement on mRNA and subsequently may affect the efficiency of co-translational (in vivo) protein folding1,7,10,11. To understand the process of protein folding in vivo, in the cell, that is ultimately coupled to the process of protein synthesis it is essential to gain comprehensive insights into the impact of codon usage/tRNA content on the movement of ribosomes along mRNA during translational elongation.
Here we describe a simple technique that can be used to locate major translation pause sites for a given mRNA translated in various cell-free systems6-8. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA. The rationale is that at low-frequency codons, the increase in the residence time of the ribosomes results in increased amounts of nascent peptides of the corresponding sizes. In vitro transcribed mRNA is used for in vitro translational reactions in the presence of radioactively labeled amino acids to allow the detection of the nascent chains. In order to isolate ribosome bound nascent polypeptide complexes the translation reaction is layered on top of 30% glycerol solution followed by centrifugation. Nascent polypeptides in polysomal pellet are further treated with ribonuclease A and resolved by SDS PAGE. This technique can be potentially used for any protein and allows analysis of ribosome movement along mRNA and the detection of the major pause sites. Additionally, this protocol can be adapted to study factors and conditions that can alter ribosome movement and thus potentially can also alter the function/conformation of the protein.

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

A technique to identify translational pause sites on mRNA is described. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA, followed by the size analysis of the nascent chains using a denaturing gel electrophoresis.

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the ...

Production of Xenopus tropicalis Egg Extracts to Identify Microtubule-associated RNAs

Many organisms localize mRNAs to specific subcellular destinations to spatially and temporally control gene expression. Recent studies have demonstrated that the majority of the transcriptome is localized to a nonrandom position in cells and embryos. One approach to identify localized mRNAs is to biochemically purify a cellular structure of interest and to identify all associated transcripts. Using recently developed high-throughput sequencing technologies it is now straightforward to identify all RNAs associated with a subcellular structure. To facilitate transcript identification it is necessary to work with an organism with a fully sequenced genome. One attractive system for the biochemical purification of subcellular structures are egg extracts produced from the frog Xenopus laevis. However, X. laevis currently does not have a fully sequenced genome, which hampers transcript identification. In this article we describe a method to produce egg extracts from a related frog, X. tropicalis, that has a fully sequenced genome. We provide details for microtubule polymerization, purification and transcript isolation. While this article describes a specific method for identification of microtubule-associated transcripts, we believe that it will be easily applied to other subcellular structures and will provide a powerful method for identification of localized RNAs.

Production of Xenopus tropicalis Egg Extracts to Identify Microtubule-associated RNAs

We describe the collection of unfertilized Xenopus tropicalis eggs and production of a meiosis II-arrested egg extract. This egg extract can be used to purify microtubules and microtubule-associated RNAs.

Many  organisms localize mRNAs to specific subcellular destinations to spatially and  temporally control gene expression. Recent studies have demonstrated ...

Dithranol as a Matrix for Matrix Assisted Laser Desorption/Ionization Imaging on a Fourier Transform Ion Cyclotron Resonance Mass Spectrometer

Mass spectrometry imaging (MSI) determines the spatial localization and distribution patterns of compounds on the surface of a tissue section, mainly using MALDI (matrix&#160;assisted laser desorption/ionization)-based analytical techniques. New matrices for small-molecule MSI, which can improve the analysis of low-molecular weight (MW) compounds, are needed. These matrices should provide increased analyte signals while decreasing MALDI background signals. In addition, the use of ultrahigh-resolution instruments, such as Fourier transform ion cyclotron resonance (FTICR) mass spectrometers, has&#160;the ability to resolve analyte signals from matrix signals, and this can partially overcome many problems associated with the background originating from the MALDI matrix. The reduction in the intensities of the metastable matrix clusters by FTICR MS can also help to overcome some of the interferences associated with matrix peaks on other instruments. High-resolution instruments such as the FTICR mass spectrometers are advantageous as they can produce distribution patterns of many compounds simultaneously while still providing confidence in chemical identifications. Dithranol (DT; 1,8-dihydroxy-9,10-dihydroanthracen-9-one) has previously been reported as a MALDI matrix for tissue imaging. In this work, a protocol for the use of DT for MALDI imaging of endogenous lipids from the surfaces of mammalian tissue sections, by positive-ion MALDI-MS, on an ultrahigh-resolution hybrid quadrupole FTICR instrument has been provided.

Dithranol (DT; 1,8-dihydroxy-9,10-dihydroanthracen-9-one) has previously been reported as a MALDI matrix for tissue imaging of small molecules; protocols for the use of DT for the MALDI imaging of endogenous lipids on the surface of tissue sections&#160;by positive-ion MALDI-MS on an ultrahigh-resolution quadrupole-FTICR instrument are provided here.

Mass spectrometry imaging (MSI) determines the spatial localization and distribution patterns of compounds on the surface of a tissue section, mainly ...

Capture Compound Mass Spectrometry - A Powerful Tool to Identify Novel c-di-GMP Effector Proteins

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis (diguanylate cyclases) and degradation (phosphodiesterases) of the second messenger c-di-GMP. In contrast, little information is available regarding the molecular mechanisms and cellular components through which this signaling molecule regulates a diverse range of cellular processes. Most of the known effector proteins belong to the PilZ family or are degenerated diguanylate cyclases or phosphodiesterases that have given up on catalysis and have adopted effector function. Thus, to better define the cellular c-di-GMP network in a wide range of bacteria experimental methods are required to identify and validate novel effectors for which reliable in silico predictions fail.
We have recently developed a novel Capture Compound Mass Spectrometry (CCMS) based technology as a powerful tool to biochemically identify and characterize c-di-GMP binding proteins. This technique has previously been reported to be applicable to a wide range of organisms1. Here we give a detailed description of the protocol that we utilize to probe such signaling components. As an example, we use Pseudomonas aeruginosa, an opportunistic pathogen in which c-di-GMP plays a critical role in virulence and biofilm control. CCMS identified 74% (38/51) of the known or predicted components of the c-di-GMP network. This study explains the CCMS procedure in detail, and establishes it as a powerful and versatile tool to identify novel components involved in small molecule signaling.

The ubiquitous second messenger c-di-GMP controls growth and behavior of many bacteria. We have developed a novel Capture Compound Mass Spectrometry based technology to biochemically identify and characterize c-di-GMP binding proteins in virtually any bacterial species.

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Bottom-up and Shotgun Proteomics to Identify a Comprehensive Cochlear Proteome