We would like to acknowledge the UF-ICBR proteomics facility for their processing of our samples. This work was supported in part by the National Institutes of Health R01 HL136759-01A1 (CAP).

The Institutional Animal Care and Use Committee from University of Florida approved all animal studies.
1. Preparation of reagents
<ol>
	<li>Prepare CHAPS Lysis Buffer (150 mM KCl, 50 mM HEPES pH = 7.4, 0.1% CHAPS, and 1 protease inhibitor cocktail tablet per 50 mL of buffer). Buffer without protease inhibitors can be stored at 4 &#176;C for up to 6 months or buffer with protease inhibitor stored at -20 &#176;C up to 1 year.</li>
	<li>Prepare 100 mM triethylammonium bicarbonate (TEAB): Add 500 &#181;L of 1 M TEAB to 4.5 mL of ultrapure water.</li>
	<li>Prepare 200 mM tris (2-carboxyethyl) phosphine hydrochloride (TCEP): Add 70 &#181;L of 0.5 M TCEP, a denaturing reagent, to 70 &#181;L of ultrapure water. Then add 35 &#181;L of the 1 M TEAB.</li>
	<li>Prepare 5% hydroxylamine: Add 50 &#181;L of 50% hydroxylamine to 450 &#181;L of 100 mM TEAB.</li>
</ol>
2. Protein extraction
<ol>
	<li>Isolate quadriceps muscle from a euthanized mouse according to an IACUC approved protocol. Freeze and maintain at -80 &#176;C or continue with the protocol for immediate use.</li>
	<li>Cut to isolate approximately 10 mg of fresh or frozen quadriceps mouse tissue. Separate fibers using tweezers when working with skeletal muscle. Alternatively, if working with cell cultures, resuspend ~3 x 106 cells in 300 &#181;L of CHAPS lysis buffer and skip to step 2.4.</li>
	<li>Homogenize tissue using a bead disrupter using 2 mL tubes filled with approximately 200 &#181;L of 1 mm zirconia/silica beads and 500 &#181;L of CHAPS lysis buffer. Scale up or down as appropriate (e.g., 5 mg of tissue in 250 &#181;L of CHAPS lysis buffer).</li>
	<li>Perform sonication (10x for 10 s each with 50% amplitude and 30 s intervals on ice) to release protein bound to DNA. The same DNA degradation results can be achieved either with syringe lysis by passing the lysate 10x through a 21 G needle attached to a 1 mL syringe, or by benzonase incubation (44 U/mL) at 37 &#176;C for 30 min.</li>
	<li>Centrifuge the lysate at 16,000 x g for 10 min at 4 &#176;C and transfer the supernatant to a new centrifuge tube.</li>
</ol>
3. Protein measurement
<ol>
	<li>Determine the protein concentration of the supernatant using established protocols (see Table of Materials). 
	NOTE: It is best to use samples at &#8805;2 &#181;g/&#181;L, but less concentrated samples may also be used. If a less concentrated sample is used it will be necessary to appropriately adjust the volumes of the reducing/alkylating reagents in step 5.1.</li>
	<li>Prepare a BSA standard curve dilution using CHAPS lysis buffer.</li>
	<li>Follow the manufacturer&#39;s instructions, and after 15 min, read the absorbance at 750 nm.</li>
</ol>
4. Reducing/alkylating reagent treatment
<ol>
	<li>Transfer 200 &#181;g of protein per condition into a new centrifuge tube and adjust to a final volume of 100 &#181;L using CHAPS lysis buffer. It is possible to scale up to 200 &#181;L when the protein concentration is too low, but do not forget to appropriately adjust the volume of reducing/alkylating reagent.</li>
	<li>Add 5 &#181;L of the 200 mM TCEP and incubate samples at 55 &#176;C for 1 h.</li>
	<li>Immediately prior to use, prepare 375 mM iodoacetamide by dissolving one tube of iodoacetamide (i.e., 9 mg) into 132 &#181;L of 100 mM TEAB. Protect this solution from light.</li>
	<li>Add 5 &#181;L of 375 mM iodoacetamide to the sample and incubate for 30 min at room temperature (RT) protected from light.</li>
</ol>
5. Methanol/chloroform precipitation20
<ol>
	<li>Add 400 &#181;L of methanol to each 100 &#181;L of protein and briefly vortex samples.</li>
	<li>Centrifuge at 9,000 x g for 10 s at RT. This is to incorporate liquids deposited on the sides of the sample tube.</li>
	<li>Add 100 &#181;L of chloroform to the mixture and briefly vortex. Use 200 &#181;L of chloroform if the sample has a high concentration of phospholipids.</li>
	<li>Centrifuge at 9,000 x g for 10 s at RT. This is to incorporate liquids deposited on the sides of the sample tube.</li>
	<li>Add 300 &#181;L of water and vortex vigorously. It is important to obtain a homogenous solution.</li>
	<li>Centrifuge at 9,000 x g for 1 min at RT. Be extremely careful to avoid disturbing the layers when transferring the tube to a rack. 
	NOTE: The tube should now contain three phases: 1) Top layer (i.e., supernatant), a mixture of water and methanol; 2) Middle layer (i.e., interphase), white precipitated protein; and 3) Bottom layer (i.e., bottom phase), chloroform.</li>
	<li>Carefully remove the supernatant.</li>
	<li>Add 300 &#181;L of methanol to the remaining interphase and bottom phase. Vortex vigorously.</li>
	<li>Centrifuge at 9,000 x g for 2 min at RT. Be extremely careful to avoid disturbing the layers when transferring the tube to a rack.</li>
	<li>Carefully remove the supernatant.</li>
	<li>Gently aspirate as much liquid as possible under a stream of air (e.g., using a vacuum concentrator) at RT until the pellet is just a bit moist (~10 min). As the necessary time may be different for each sample, check every 2 min to assess. Store the pellet at -80 &#176;C until further processing.</li>
</ol>
6. Protein digestion
<ol>
	<li>Resuspend the precipitated protein pellet in 100 &#181;L of TEAB lysis buffer. 
	NOTE: It is optional to measure the protein concentration at this step.</li>
	<li>Immediately prior to use, prepare 1 &#181;g/&#181;L trypsin by adding 100 &#181;L of the trypsin storage solution (50 mM acetic acid) to the bottom of the 100 &#181;g trypsin glass vial and incubate for 5 min at RT. Store remaining reagent in single-use doses at -80 &#176;C.</li>
	<li>Add 2.5 &#181;L of trypsin per 100 &#181;g of protein. Digest the sample overnight at 37 &#176;C. This step is crucial for complete solubilization of the protein; do not modify these conditions. Following the digestion, it is optional to measure the protein concentration using standard protein assays.</li>
</ol>
7. Peptide labeling
<ol>
	<li>Immediately prior to use, equilibrate the TMT label kit reagents to RT.</li>
	<li>Dissolve each of the 0.8 mg TMT tag vials through the addition of 41 &#181;L of anhydrous acetonitrile to each tube. Incubate the reagent for 5 min at RT with occasional vortexing. Briefly centrifuge the tubes. 
	NOTE: A concentration of 0.8 mg of TMT tag is usually enough to label two sets. Other investigators, however, have demonstrated that this concentration can be reduced further and still yield reliable data15.</li>
	<li>Carefully add 41 &#181;L of the TMT label reagent to each 100 &#181;L sample.</li>
	<li>Incubate the reaction for 1 h at RT.</li>
	<li>Add 8 &#181;L of 5% hydroxylamine to the sample and incubate for 15 min to quench the reaction.</li>
	<li>Split the samples into equal amounts in a new centrifuge tube and store at -80 &#176;C. 
	NOTE: In this step the samples are stable and can be submitted for mass spectroscopy. It is optional to measure concentration at this point using standard protein assays.</li>
</ol>
8. Mass spectroscopy
<ol>
	<li>Submit the samples to a proteomics facility (this study used the UF ICBR Proteomics Core facility), where all samples are combined and purified using C18 spin columns. 
	NOTE: Discuss with the core facility how to submit the samples prior to preparing them to confirm the exact steps they prefer for submissions.</li>
	<li>Request the following procedures per combined multiplex sample: solid phase extraction, HPLC (SCX, SE), zip tip, and LC-MS/MS (2 h gradient for protein ID, if &#62;10QE Plus).</li>
	<li>Once the data are collected, the core facility will process RAW files using vendor-supplied software for protein identification.</li>
</ol>
9. Data analysis
<ol>
	<li>Data are typically delivered from the core back to the user in the 7z format, which can require approximately 16 GB of disk space per each dataset (in this case 11 samples). For data processing, make sure a computer is available that is at least 3.4 GHz.</li>
	<li>Extract files using 7-Zip File Manager. These extracted files contain RAW data, pdStudy format file, and pdResultView format file. Save all files for further analyses.</li>
	<li>Open file using Proteome Discovery 2.2 Software. 
	NOTE: The file format is &#34;File name.pdStudy&#34;. If the &#34;File name.pdResultView&#34; is opened it is not possible to select the control sample.</li>
	<li>Select control samples on &#34;Samples&#34; panel.</li>
	<li>Open Result by selecting ID on &#34;Analysis Results&#34; panel.</li>
	<li>Export to spreadsheet software.</li>
	<li>Save raw data (all proteins identified).</li>
	<li>Open the spreadsheet software file. This will contain all proteins that have been identified.</li>
	<li>In the spreadsheet software file use the &#34;Filter&#34; function to screen &#34;Protein FDR Confidence: Combined&#34; in high (column B), &#34;#Unique Peptides&#34; higher than 2 (column K), and either one of &#34;Abundance Ratio&#34; blank exclusively (column S up to W).</li>
	<li>Insert a column for &#34;p-value&#34; calculation with the function 
	=TTEST(control group,experimental group, tails, type)</li>
	<li>Insert a column for &#34;Statistical Significance&#34; with the function 
	=IF(p-value&#60;0.05, &#34;Significance&#34;,&#34;NS&#34;)</li>
	<li>Use the &#34;Filter&#34; function to screen &#34;Statistical Significance&#34; showing &#34;Significance&#34;. The result shows the analyzed proteins with the statistical significance in the control group and experimental group.</li>
	<li>Determine significantly higher or lower protein expression abundance in the experimental group compared to the control group, insert a column for &#34;Regulation&#34; with the function 
	=IF(AVERAGE(controlgroup)&#62;AVERAGE(experimentalgroup),&#34;Upregulated&#34;,&#34;Downregulated&#34;)</li>
</ol>
10. Methods to evaluate significant hits
<ol>
	<li>To identify protein-protein interactions between the significant hits identified in the TMT studies, use Search Tool for the Retrieval of Interacting Genes/Proteins (STRING) version 11.021: <a href="https://string-db.org/" target="_blank">https://string-db.org/</a></li>
	<li>To classify by groups (i.e., molecular function, biological processes, and protein classes) use Protein Analysis through Evolutionary Relationships (PANTHER) ontology classification software22: <a href="http://www.pantherdb.org/" target="_blank">http://www.pantherdb.org/</a></li>
	<li>To identify protein interactions in a variety of pathways, use pathway analysis software23.</li>
</ol>
11. Proteomic data upload to a repository bank
<ol>
	<li>To submit proteomic data to the Proteomics IDEntificantions Database (PRIDE) or Mass Spectrometry Interactive Virtual Environment (MassIVE) include the following information: the peak list files (processed mass spectrum files in a standard format such as mzXML, mzML, or MGF), result files (spectrum identifications in a standard format such as mzIdentML or mzTab), and raw spectrum files (raw mass spectrum files in a nonstandard or instrument-specific format such as .RAW files or .WIFF files).</li>
	<li>To submit, create an account and include information such as affiliation and project details. Then, select the files listed in step 11.1 and upload them.</li>
	<li>To create an official dataset, run a submission workflow on these uploaded files. 
	NOTE: Following submission, the dataset will be private in the repository bank. With the private option, the data are only available to authorized users. There are two additional options: 1) Shared dataset, which gives access to journal reviewers and collaborators; or 2) Public dataset, which will show up in public dataset searches. Another important feature of these repositories is the ability to update the uploaded data and associate subsequent publications with the existing dataset.</li>
</ol>

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The authors have nothing to disclose.

To successfully prepare samples for proteomic analysis using TMT-based isobaric stable isotope labeling methodologies, it is crucial to perform protein extractions very carefully at 4 &#176;C and to use a lysis buffer that contains a protease inhibitor cocktail24,25. The protease inhibitor cocktail is a crucial reagent to avoid unexpected protein degradation during protein digestion. One key difference between our protocol and the current one provided by the vendor is that we strongly recommend the use of CHAPS lysis buffer based upon our experience with mammalian cells and tissues. We also suggest using a methanol/chloroform protein precipitation approach for both cell pellets and tissues.
Ideally, protein extraction, measurement, reducing/alkylating reagent treatments, and methanol/chloroform precipitations are all performed on the same day. Following this recommendation will result in more accurate protein concentrations for subsequent labeling. The protein precipitation step is important for the removal of reagents that will interfere with tandem mass spectrometry. Including the precipitation step significantly enhances the resolution of TMT26. In sum, the major advantages of our TMT protocol are the high labeling efficiencies for different types of samples, its reproducibility, and the reliable data acquired.
As the multiplex nature of this TMT untargeted proteomics strategy continues to expand, it will progressively enhance the ability of researchers across a wide variety of fields to make novel discoveries. Specifically in the biomedical field, we and others have found this technology increasingly informative in studies exploring novel mechanisms of action in disease and relative impacts of various therapeutics. For all of these reasons, this powerful technology complements the repertoire of other OMICS approaches used in modern research studies and provides key information that can guide further therapeutic development.

The term &#34;proteomics&#34; was first defined as the large-scale characterization of the entire protein complement of a cell, tissue, or organism1. Proteomic analyses enable the investigation of mechanisms and cellular processes involved in disease development, therapeutic pathways, and healthy systems using techniques to perform relative quantitation of protein expression levels2. The initial descriptions of such studies were published in 1975 and demonstrated the use of two-dimensional polyacrylamide gel electrophoresis (2D-PAGE) for this purpose1,3. The 2D method separates proteins based on charge (isoelectric focusing, IEF) and molecular mass (sodium dodecyl sulfate polyacrylamide gel electrophoresis, or SDS-PAGE)4. For years, the combination of 2D-PAGE and subsequent tandem mass spectrometry performed on each gel component was the most common untargeted protein expression analysis technique performed and identified numerous previously unknown protein expression profiles5,6. General disadvantages to the 2D-PAGE approach are that it is time-consuming, does not work well for hydrophobic proteins, and there are limitations in the total number of proteins assessed due to low sensitivity7,8.
The stable isotope labeling by amino acids in cell culture (SILAC) method became the next popular approach to identify and quantify protein abundance in samples9. It consists of metabolic labeling of cells that are incubated in medium lacking a standard essential amino acid and supplemented with an isotope-labeled version of that specific amino acid10. The advantage of this technique is its efficiency and precise labeling9. The main limitation to the SILAC approach is primarily the reduced cell growth rate caused by isotope label incorporation, which can be particularly challenging in relatively sensitive cell lines modeling human diseases11.
In 2003, a novel and robust proteomics technique involving tandem mass tag (TMT) isobaric labels was introduced to the field12. TMT labeling is a powerful method due to its increased sensitivity to detect relative protein expression levels and posttranslational modifications13. As of this publication date, TMT kits have been developed that can simultaneously label 6, 10, 11, or 16 samples. As a result, it is possible to measure peptide abundance in multiple conditions with biological replicates at the same time14,15,16. We recently used TMT to characterize the cardiac proteomic profile of a mouse model of Barth Syndrome (BTHS)17. In so doing, we were able to demonstrate widespread improvement in the cardiac profiles of BTHS mice treated with gene therapy and identify novel proteins impacted by BTHS that revealed novel therapeutic pathways involved in cardiomyopathies.
Here, we describe a detailed method to perform multiplex TMT quantitative proteomics analyses using tissue samples or cell pellets. It can be beneficial to perform the sample preparation and labeling prior to submission to a core because the labeled tryptic peptides are more stable than raw frozen samples, not all cores have experience handling all sample types, and preparing samples in a laboratory can save time for cores, which often have long backlogs. For detailed descriptions of the mass spectroscopy portion of this process please see Kirshenbaum et al. and Perumal et al.18,19.
The sample preparation protocol consists of the following major steps: extraction, quantification, precipitation, digestion, and labeling. The major benefits of this optimized protocol are that it reduces the costs of labeling, improves protein extraction, and consistently generates high-quality data. In addition, we describe how to analyze TMT data to screen thousands of proteins in a short amount of time. We hope that this protocol encourages other research groups to consider incorporating this powerful methodology into their studies.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 M Triethylammonium bicarbonate (TEAB), 50 mL</td>
			<td>Thermo Fisher</td>
			<td>90114</td>
			<td>Reagent for protein labeling</td>
		</tr>
		<tr>
			<td>50% Hydroxylamine, 5 mL</td>
			<td>Thermo Fisher</td>
			<td>90115</td>
			<td>Reagent for protein labeling</td>
		</tr>
		<tr>
			<td>Acetic acid</td>
			<td>Sigma</td>
			<td>A6283</td>
			<td>Reagent for protein digestion</td>
		</tr>
		<tr>
			<td>Anhydrous acetonitrile, LC-MS Grade</td>
			<td>Thermo Fisher</td>
			<td>51101</td>
			<td>Reagent for protein labeling</td>
		</tr>
		<tr>
			<td>Benzonaze nuclease</td>
			<td>Sigma-Aldrich</td>
			<td>E1014</td>
			<td>DNA shearing</td>
		</tr>
		<tr>
			<td>Bond-Breaker TCEP solution, 5 mL</td>
			<td>Thermo Fisher</td>
			<td>77720</td>
			<td>Reagent for protein labeling</td>
		</tr>
		<tr>
			<td>BSA standard</td>
			<td>Thermo</td>
			<td>23209</td>
			<td>Reagent for protein measurement</td>
		</tr>
		<tr>
			<td>CHAPS</td>
			<td>Thermo Fisher</td>
			<td>28300</td>
			<td>Reagent for protein extraction</td>
		</tr>
		<tr>
			<td>Chloroform</td>
			<td>Fisher</td>
			<td>BP1145-1</td>
			<td>Reagent for protein precipitation</td>
		</tr>
		<tr>
			<td>cOmplete, EDTA-free Protease Inhibitor Cocktail Tablet</td>
			<td>Roche</td>
			<td>4693132001</td>
			<td>Reagent for protein extraction</td>
		</tr>
		<tr>
			<td>DC Protein Assay</td>
			<td>BioRad</td>
			<td>500-0116</td>
			<td>Reagent for protein measurement</td>
		</tr>
		<tr>
			<td>Excel</td>
			<td>Microsoft Office</td>
			<td></td>
			<td>Software for data analyses</td>
		</tr>
		<tr>
			<td>Heat block</td>
			<td>VWR analog</td>
			<td>12621-104</td>
			<td>Equipment for protein digestion incubation</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>RDD002</td>
			<td>Reagent for protein extraction</td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Fisher</td>
			<td>A452-4</td>
			<td>Reagent for protein precipitation</td>
		</tr>
		<tr>
			<td>Pierce Trypsin Protease, MS Grade</td>
			<td>Thermo Fisher</td>
			<td>90058</td>
			<td>Reagent for protein digestion</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma</td>
			<td>46436</td>
			<td>Reagent for protein extraction</td>
		</tr>
		<tr>
			<td>Sigma Plot 14.0</td>
			<td>Sigma Plot 14.0</td>
			<td></td>
			<td>Software for data analyses</td>
		</tr>
		<tr>
			<td>Sonicator</td>
			<td>Fisher Scientific</td>
			<td>FB120</td>
			<td>DNA shearing</td>
		</tr>
		<tr>
			<td>Spectra Max i3x Multi-Mode Detection Platform</td>
			<td>Molecular Devices</td>
			<td></td>
			<td>Plate reader for protein measurement</td>
		</tr>
		<tr>
			<td>Thermo Scientific Pierce Quantitative Colorimetric Peptide Assay</td>
			<td>Thermo Fisher</td>
			<td>23275</td>
			<td>Reagent for protein measurement</td>
		</tr>
		<tr>
			<td>Thermo Scientific Pierce Quantitative Fluorescent Peptide Assay</td>
			<td>Thermo Fisher</td>
			<td>23290</td>
			<td>Reagent for protein measurement</td>
		</tr>
		<tr>
			<td>Thermo Scientific Proteome Discoverer Software</td>
			<td>Thermo Fisher</td>
			<td>OPTON-30945</td>
			<td>Software for data analyses</td>
		</tr>
		<tr>
			<td>TMT 10plex Isobaric Label Reagent Set 0.8 mg, sufficient reagents for one 10plex isobaric experiment</td>
			<td>Thermo Fisher</td>
			<td>90110</td>
			<td>Reagent for protein labeling</td>
		</tr>
		<tr>
			<td>TMT11-131C Label Reagent 5 mg</td>
			<td>Thermo Fisher</td>
			<td>A34807</td>
			<td>Reagent for protein labeling</td>
		</tr>
		<tr>
			<td>Water, LC-MS Grade</td>
			<td>Thermo Fisher</td>
			<td>51140</td>
			<td>Reagent for protein extraction</td>
		</tr>
	</tbody>
</table>

tmt sample preparation for proteomics facility submission and subsequent data analysis

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view of the impact of a disease, treatment, or other condition on the proteome as a whole. This report provides a detailed protocol for the extraction, quantification, precipitation, digestion, labeling, and subsequent data analysis of protein samples. Our optimized TMT labeling protocol requires a lower tag-label concentration and achieves consistently reliable data. We have used this protocol to evaluate protein expression profiles in a variety of mouse tissues (i.e., heart, skeletal muscle, and brain) as well as cells cultured in vitro. In addition, we demonstrate how to evaluate thousands of proteins from the resulting dataset.

Healthy and diseased cells were lysed in CHAPS buffer, prepared as detailed in our TMT labeling method, and submitted to the University of Florida Interdisciplinary Center for Biotechnology Research (UF-ICBR) Proteomics Core for Liquid Chromatography with tandem mass spectrometry. Following data acquisition and delivery from the core, the dataset was opened in vendor-supplied software and the following cutoff filters were applied: &#8805;2 unique peptides, reporter ions for each protein sample present in all channels, and include only significantly altered proteins (p &#8804; 0.05). Table 1 summarizes the data: 39,653 total peptides, of which 7,211 have equal or greater than two unique peptides, and 3,829 include reporter ions for all channels. The p values for these 3,829 peptides were calculated by Student&#39;s t test and p &#8804; 0.05 was considered significant. In addition, a fold-change cutoff was used to determine the relative distribution of proteins from diseased compared to healthy cells: downregulated (blue) or upregulated (red) (Figure 1).
The list of significantly dysregulated protein expression was assessed using the PANTHER ontology classification system and STRING analyses. Panther analyses showed a categorized list of proteins based upon significantly lower (Figure 2A) or higher abundance in diseased cells based upon molecular function (Figure 2B). String analyses of proteins of significantly lower (Figure 2C) and higher (Figure 2D) abundance identified multiple interactions and strong associations between proteins.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60970/60970fig1.jpg" /> 
Figure 1: Volcano plot displaying proteins whose abundance was not significantly altered (black), significantly lowered (blue), or significantly increased (red) in diseased vs. healthy control cells. <a href="https://www.jove.com/files/ftp_upload/60970/60970fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60970/60970fig2.jpg" /> 
Figure 2: Representative evaluations of significantly dysregulated hits identified by PANTHER (A, B) and String (C, D) of significantly lower or higher abundance proteins. <a href="https://www.jove.com/files/ftp_upload/60970/60970fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Total peptides</td>
			<td>Total identified</td>
			<td>&#8805; 2 unique peptides</td>
			<td>Quantified proteins</td>
			<td colspan="2">Significantly altered proteins</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td>Low</td>
			<td>High</td>
		</tr>
		<tr>
			<td>39653</td>
			<td>7211</td>
			<td>4457</td>
			<td>3829</td>
			<td>296</td>
			<td>108</td>
		</tr>
	</tbody>
</table>
Table 1: Representative table of quantified proteins per dataset analysis.

Watch this Scientific Journal Video about TMT Sample Preparation for Proteomics Facility Submission and Subsequent Data Analysis at JoVE.com

TMT Sample Preparation for Proteomics Facility Submission and Subsequent Data Analysis

We present an optimized tandem mass tag (TMT) labeling protocol that includes detailed information for each of the following steps: protein extraction, quantification, precipitation, digestion, labeling, submission to a proteomics facility, and data analyses.

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view ...

tmt-sample-preparation-for-proteomics-facility-submission-subsequent

Research

JoVE Journal

Biochemistry

12.5K Views.  University of Florida. We present an optimized tandem mass tag (TMT) labeling protocol that includes detailed information for each of the following steps: protein extraction, quantification, precipitation, digestion, labeling, submission to a proteomics facility, and data analyses.

Video: TMT Sample Preparation for Proteomics Facility Submission and Subsequent Data Analysis

Lipid Droplet Isolation for Quantitative Mass Spectrometry Analysis

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of virion morphogenesis. Quantitative lipid droplet proteome analysis can be used to identify proteins that localize to or are displaced from lipid droplets under conditions such as virus infections. Here, we describe a protocol that has been successfully used to characterize the changes in the lipid droplet proteome following infection with HCV. We use Stable Isotope Labeling with Amino Acids in Cell Culture (SILAC) and thus label the complete proteome of one population of cells with &#34;heavy&#34; amino acids to quantitate the proteins by mass spectrometry. For lipid droplet isolation, the two cell populations (i.e.&#160;HCV-infected/&#34;light&#34; amino acids and uninfected control/&#34;heavy&#34; amino acids) are mixed 1:1 and lysed mechanically in hypotonic buffer. After removing the nuclei and cell debris by low speed centrifugation, lipid droplet-associated proteins are enriched by two subsequent ultracentrifugation steps followed by three washing steps in isotonic buffer. The purity of the lipid droplet fractions is analyzed by western blotting with antibodies recognizing different subcellular compartments. Lipid droplet-associated proteins are then separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coomassie staining. After tryptic digest, the peptides are quantified by liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS). Using this method, we identified proteins recruited to lipid droplets upon HCV infection that might represent pro- or antiviral host factors. Our method can be applied to a variety of different cells and culture conditions, such as infection with pathogens, environmental stress, or drug treatment.

Lipid droplets are important organelles for the replication of several pathogens, including the Hepatitis C Virus (HCV). We describe a method to isolate lipid droplets for quantitative mass spectrometry of associated proteins; it can be used under a variety of conditions, such as virus infection, environmental stress, or drug treatment.

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of ...

Optimal Preparation of Formalin Fixed Samples for Peptide Based Matrix Assisted Laser Desorption/Ionization Mass Spectrometry Imaging Workflows

The use of matrix-assisted laser desorption/ionization, mass spectrometry imaging (MALDI MSI) has rapidly expanded, since this technique analyzes a host of biomolecules from drugs and lipids to N-glycans. Although various sample preparation techniques exist, detecting peptides from formaldehyde preserved tissues remains one of the most difficult challenges for this type of mass spectrometric analysis. For this reason, we have created and optimized a robust methodology that preserves the spatial information contained within the sample, while eliciting the greatest number of ionizable peptides. We have also aimed to achieve this in a cost effective and simple way, thereby eliminating potential bias or preparation error, which can occur when using automated instrumentation. The end result is a reproducible and inexpensive protocol.

This protocol describes a reproducible and reliable method for the sublimation-based preparation of formalin fixed tissue destined for imaging mass spectrometry.

The use of matrix-assisted laser desorption/ionization, mass spectrometry imaging (MALDI MSI) has rapidly expanded, since this technique analyzes a host ...

Preparation of Chloroplast Sub-compartments from Arabidopsis for the Analysis of Protein Localization by Immunoblotting or Proteomics

Chloroplasts are major components of plant cells. Such plastids fulfill many crucial functions, such as assimilation of carbon, sulfur and nitrogen as well as synthesis of essential metabolites. These organelles consist of the following three key sub-compartments. The envelope, characterized by two membranes, surrounds the organelle and controls the communication of the plastid with other cell compartments. The stroma is the soluble phase of the chloroplast and the main site where carbon dioxide is converted into carbohydrates. The thylakoid membrane is the internal membrane network consisting of grana (flat compressed sacs) and lamellae (less dense structures), where oxygenic photosynthesis takes place. The present protocol describes step by step procedures required for the purification, using differential centrifugations and Percoll gradients, of intact chloroplasts from Arabidopsis, and their fractionation, using sucrose gradients, in three sub-compartments (i.e., envelope, stroma, and thylakoids). This protocol also provides instructions on how to assess the purity of these fractions using markers associated to the various chloroplast sub-compartments. The method described here is valuable for subplastidial localization of proteins using immunoblotting, but also for subcellular and subplastidial proteomics and other studies.

Here, we describe a method to purify intact chloroplasts from Arabidopsis leaves and their three main sub-compartments (envelope, stroma, and thylakoids), using a combination of differential centrifugations, continuous Percoll gradients, and discontinuous sucrose gradients. The method is valuable for subplastidial and subcellular localization of proteins by immunoblotting and proteomics.

Chloroplasts are major components of plant cells. Such plastids fulfill many crucial functions, such as assimilation of carbon, sulfur and nitrogen as ...

Fully Autonomous Characterization and Data Collection from Crystals of Biological Macromolecules

High-brilliance X-ray beams coupled with automation have&#160;led to the use of synchrotron-based macromolecular X-ray crystallography (MX) beamlines for even the most challenging projects in structural biology. However, most facilities still require the presence of a scientist on site to perform the experiments. A new generation of automated beamlines dedicated to the fully automatic characterization of, and data collection from, crystals of biological macromolecules has recently been developed. These beamlines represent a new tool for structural biologists to screen the results of initial crystallization trials and/or the collection of large numbers of diffraction data sets, without users having to control the beamline themselves. Here we show how to set up an experiment for automatic screening and data collection, how an experiment is performed at the beamline, how the resulting data sets are processed, and how, when possible, the crystal structure of the biological macromolecule is solved.

Here, we describe how to use the automated screening and data collection options available at some synchrotron beamlines. Scientists send cryocooled samples to the synchrotron, and the diffraction properties are screened, the data sets are collected and processed and, where possible, a structure solution is carried out&#8212;all without human intervention.

High-brilliance X-ray beams coupled with automation have&#160;led to the use of synchrotron-based macromolecular X-ray crystallography (MX) beamlines for ...

Sample Preparation for Probe Electrospray Ionization Mass Spectrometry

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ratios (m/z), which are useful to deduce molecular weights and structures. While it is essentially a destructive analytical method, recent advancements in the ambient ionization technique have enabled us to acquire data while leaving tissue in a relatively intact state in terms of integrity. Probe electrospray ionization (PESI) is a so-called direct method because it does not require complex and time-consuming pretreatment of samples. A fine needle serves as a sample picker, as well as an ionization emitter. Based on the very sharp and fine property of the probe tip, destruction of the samples is minimal, allowing us to acquire the real-time molecular information from living things in situ. Herein, we introduce three applications of PESI-MS technique that will be useful for biomedical research and development. One involves the application to solid tissue, which is the basic application of this technique for the medical diagnosis. As this technique requires only 10 mg of the sample, it may be very useful in the routine clinical settings. The second application is for in vitro medical diagnostics where human blood serum is measured. The ability to measure fluid samples is also valuable in various biological experiments where a sufficient volume of sample for conventional analytical techniques cannot be provided. The third application leans toward the direct application of probe needles in living animals, where we can obtain real-time dynamics of metabolites or drugs in specific organs. In each application, we can infer the molecules that have been detected by MS or use artificial intelligence to obtain a medical diagnosis.

This article introduces sample preparation methods for a unique real-time analytical method based on the ambient mass spectrometry. This method lets us perform real-time analysis of the biological molecules in vivo without any special pretreatments.

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ...

Analysis of SEC-SAXS data via EFA deconvolution and Scatter

BioSAXS is a popular technique used in molecular and structural biology to determine the solution structure, particle size and shape, surface-to-volume ratio and conformational changes of macromolecules and macromolecular complexes. A high quality SAXS dataset for structural modeling must be from monodisperse, homogeneous samples and this is often only reached by a combination of inline chromatography and immediate SAXS measurement. Most commonly, size-exclusion chromatography is used to separate samples and exclude contaminants and aggregations from the particle of interest allowing SAXS measurements to be made from a well-resolved chromatographic peak of a single protein species. Still, in some cases, even inline purification is not a guarantee of monodisperse samples, either because multiple components are too close to each other in size or changes in shape induced through binding alter perceived elution time. In these cases, it may be possible to deconvolute the SAXS data of a mixture to obtain the idealized SAXS curves of individual components. Here, we show how this is achieved and the practical analysis of SEC-SAXS data is performed on ideal and difficult samples. Specifically, we show the SEC-SAXS analysis of the vaccinia E9 DNA polymerase exonuclease minus mutant.

SEC-BioSAXS measurements of biological macromolecules are a standard approach for determining solution structure of macromolecules and their complexes. Here, we analyze SEC-BioSAXS data from two types of commonly encountered SEC traces&#8212;chromatograms with fully resolved and partially resolved peaks. We demonstrate the analysis and deconvolution using scatter and BioXTAS RAW.

BioSAXS is a popular technique used in molecular and structural biology to determine the solution structure, particle size and shape, surface-to-volume ...

A Sample Preparation Pipeline for Microcrystals at the VMXm Beamline

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality have been seen for microcrystals with the development of beamline optics, beam stability and variable beam size focusing from submicron to microns, such as at the VMXm beamline at Diamond Light Source1. Further improvements in data quality will be gained through improvements in sample environment and sample preparation. Microcrystals inherently generate weaker diffraction, therefore improving the signal-to-noise is key to collecting quality X-ray diffraction data and will predominantly come from reductions in background noise. Major sources of X-ray background noise in a diffraction experiment are from their interaction with the air path before and after the sample, excess crystallization solution surrounding the sample, the presence of crystalline ice and scatter from any other beamline instrumentation or X-ray windows. The VMXm beamline comprises instrumentation and a sample preparation protocol to reduce all these sources of noise.
Firstly, an in-vacuum sample environment at VMXm removes the air path between X-ray source and sample. Next, sample preparation protocols for macromolecular crystallography at VMXm utilize a number of processes and tools adapted from cryoTEM. These include copper grids with holey carbon support films, automated blotting and plunge cooling robotics making use of liquid ethane. These tools enable the preparation of hundreds of microcrystals on a single cryoTEM grid with minimal surrounding liquid on a low-noise support. They also minimize the formation of crystalline ice from any remaining liquid surrounding the crystals. 
We present the process for preparing and assessing the quality of soluble protein microcrystals using visible light and scanning electron microscopy before mounting the samples on the VMXm beamline for X-ray diffraction experiments. We will also provide examples of good quality samples as well as those which require further optimization and strategies to do so.

The signal-to-noise ratio of data is one of the most important considerations in performing X-ray diffraction measurements from microcrystals. The VMXm beamline provides a low-noise environment and microbeam for such experiments. Here, we describe sample preparation methods for mounting and cooling microcrystals for VMXm and other microfocus macromolecular crystallography beamlines.

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality ...

Preparation of High-Temperature Sample Grids for Cryo-EM

The sample grids for cryo-electron microscopy (cryo-EM) experiments are usually prepared at a temperature optimal for the storage of biological samples, mostly at 4 &#176;C and occasionally at room temperature. Recently, we discovered that the protein structure solved at low temperature may not be functionally relevant, particularly for proteins from thermophilic archaea. A procedure was developed to prepare protein samples at higher temperatures (up to 70 &#176;C) for cryo-EM analysis. We showed that the structures from samples prepared at higher temperatures are functionally relevant and temperature dependent. Here we describe a detailed protocol for preparing sample grids at high temperature, using 55 &#176;C as an example. The experiment made use of a vitrification apparatus modified using an additional centrifuge tube, and samples were incubated at 55 &#176;C. The detailed procedures were fine-tuned to minimize vapor condensation and obtain a thin layer of ice on the grid. Examples of successful and unsuccessful experiments are provided.

This paper provides a detailed protocol for preparing sample grids at temperatures as high as 70 &#176;C, prior to plunge freezing for cryo-EM experiments.

The sample grids for cryo-electron microscopy (cryo-EM) experiments are usually prepared at a temperature optimal for the storage of biological samples, ...

Sample Preparation and Relative Quantitation using Reductive Methylation of Amines for Peptidomics Studies

Peptidomics can be defined as the qualitative and quantitative analysis of peptides in a biological sample. Its main applications include identifying the peptide biomarkers of disease or environmental stress, identifying neuropeptides, hormones, and bioactive intracellular peptides, discovering antimicrobial and nutraceutical peptides from protein hydrolysates, and can be used in studies to understand the proteolytic processes. The recent advance in sample preparation, separation methods, mass spectrometry techniques, and computational tools related to protein sequencing has contributed to the increase of the identified peptides number and peptidomes characterized. Peptidomic studies frequently analyze peptides that are naturally generated in cells. Here, a sample preparation protocol based on heat-inactivation is described, which eliminates protease activity, and extraction with mild conditions, so there is no peptide bonds cleavage. In addition, the relative quantitation of peptides using stable isotope labeling by reductive methylation of amines is also shown. This labeling method has some advantages as the reagents are commercially available, inexpensive compared to others, chemically stable, and allows the analysis of up to five samples in a single LC-MS run.

This article describes a sample preparation method based on heat-inactivation to preserve endogenous peptides avoiding degradation post-mortem, followed by relative quantitation using isotopic labeling plus LC-MS.

Peptidomics can be defined as the qualitative and quantitative analysis of peptides in a biological sample. Its main applications include identifying the ...

Sample Preparation using a Lipid Monolayer Method for Electron Crystallographic Studies

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown into highly ordered two-dimensional (2D) crystals under favorable conditions. The quality of the grown 2D crystals is crucial to the resolution of the final reconstruction via 2D image processing. Over the years, lipid monolayers have been used as a supporting layer to foster the 2D crystallization of peripheral membrane proteins as well as soluble proteins. This method can also be applied to 2D crystallization of integral membrane proteins but requires more extensive empirical investigation to determine detergent and dialysis conditions to promote partitioning to the monolayer. A lipid monolayer forms at the air-water interface such that the polar lipid head groups remain hydrated in the aqueous phase and the non-polar, acyl chains, tails partition into the air, breaking the surface tension and flattening the water surface. The charged nature or distinctive chemical moieties of the head groups provide affinity for proteins in solution, promoting binding for 2D array formation. A newly formed monolayer with the 2D array can be readily transfer into an electron microscope (EM) on a carbon-coated copper grid used to lift and support the crystalline array. In this work, we describe a lipid monolayer methodology for cryogenic electron microscopic (cryo-EM) imaging.

Lipid monolayers have been used as a foundation for forming two-dimensional (2D) protein crystals for structural studies for decades. They are stable at the air-water interface and can serve as a thin supporting material for electron imaging. Here we present the proven steps on preparing lipid monolayers for biological studies.

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown ...