Name: amide hydrogen/deuterium exchange &amp; maldi-tof mass spectrometry analysis of pak2 activation
Uploaded: 2011-11-26T14:00:00
Duration: 7 min 15 s
Description: Watch this Scientific Journal Video about Amide Hydrogen/Deuterium Exchange & MALDI-TOF Mass Spectrometry Analysis of Pak2 Activation at JoVE.com

This work is supported in part from the National Institutes of Health Grant GM-26738 (to J.A.T.).

1. Pak2 Activation
<ol>
	<li>Add the appropriate pre-tested amount of caspase 3 to 20 &mu;l of 12 mg/ml Pak2 in buffer A (50 mM Tris-HCl, 150 mM NaCl, and 2 mM DTT at pH 7.5) and incubate at 34 &deg;C for 30 min to fully cleave and activate Pak2.</li>
	<li>Analyze the cleavage products by SDS-PAGE (10 %) and Coommassie blue staining.</li>
	<li>Add the cleaved Pak2 to activation buffer B (50 mM Tris-HCl, 150 mM NaCl, 15 mM MgCl2, 10 mM ATP) at pH 7.5, in 20 &mu;l to a final concentration of 10 mg/ml Pak2, and incubate at 34 &deg;C for 30 min to fully activate Pak2 by autophosphorylation at 8 sites.</li>
	<li>Confirm full cleavage of Pak2 by migration of the p34 and p27 fragments as determined by SDS-PAGE10. Proper verification of the autophosphorylation conditions is made via autoradiography in a separate experiment using (32P)ATP or mass spectrometry to detect phosphopeptides.</li>
</ol>
2. Identification of Pepsin-Digested Pak2 Fragments
<ol>
	<li>Add 1 ml of 0.1 % Trifluroacetic Acid (TFA) at pH 2.5 to wash the agarose beads twice prior to use. Then, centrifuge the samples for 15 sec at 2,000 x g to remove the beads.</li>
	<li>Add Pak2 (20 &mu;g in 2 &mu;l of 150 mM NaCl, 50 mM Tris-HCl and 2 mM DTT at pH 7.5) to 100 &mu;l of 0.1 % TFA, and incubate the sample for 5 min with 30 &mu;l of pepsin-conjugated agarose beads.</li>
	<li>Equilibrate a reverse phase HPLC C18 column with a primary solvent system of 0.1 % TFA at pH 2.5. The pepsin digest (50 &mu;l) is eluted using a linear 100-min gradient of 0 - 80 % acetonitrile containing 0.1 % TFA at a flow rate of 0.2 ml/min. Fractions are collected every 2 min.</li>
	<li>Dry the HPLC fractions with a speed-vac (3 h) and resuspend the samples in a solution of acetonitrile (5 &mu;l) and 0.1 % TFA (5 &mu;l) at pH 2.5.</li>
	<li>Initially, each fraction is screened for peptides using the MALDI-TOF PerSeptive Biosystems Voyager DE STR (PE Biosystems, Foster City, CA, USA).</li>
	<li>The 16 fractions containing peptides are subjected to tandem mass spectrometry using a Q-TOF mass spectrometer and a QSTAR XL oMALDI MS/MS to identify the fragments.</li>
	<li>The sequence of each peptide is verified by the product ions generated by the tandem mass spectrometry with the theoretical m/z ratios based on the primary sequence of Pak2 using the Protein Prospector website (<a href="http://prospector.ucsf.edu/">http://prospector.ucsf.edu/</a>)11.</li>
</ol>
3. Measurement of Amide H/D Exchange
<ol>
	<li>Hold all solutions and reagents needed for H/D exchange at 25 &deg;C in a water bath. ATP is dissolved in water and the pH is adjusted to 7.0 prior to use. The additional 4mM ATP in the D2O buffer is to prevent ATP dissociated from Pak2 after dilution.</li>
	<li>Initiate H/D exchange by addition of 18 &mu;l of buffered D2O (50 mM MOPS pH 6.98, 125 mM NaCl) to 2 &mu;l (20 &mu;g) of Pak2 in a buffer containing 150 mM NaCl, 50 mM Tris-HCl (pH 7.5) and 2 mM DTT, resulting in a Pak2 level of 10 mg/ml and a pH of 7.0 at 0&deg;C.</li>
	<li>Incubate the samples in triplicate in the H/D exchange for 0, 0.5, 1, 3, 5, and 10 min, or 24 h.</li>
	<li>Quench the H/D exchange process by adding 180 &mu;l of ice-cold 0.1 % TFA at pH 2.2, which brings the pH to 2.5 in a final volume of 200 &mu;l. For the samples containing ATP, 180 &mu;l of ice-cold 0.11 % TFA at pH 2.2 is used to maintain the pH at 2.5 in the quenched condition.</li>
	<li>Add an aliquot (100 &mu;l) of each quenched sample to 30 &mu;l of activated pepsin-conjugated agarose beads. The pepsin digestion is allowed to proceed on ice for five min and the samples are vortexed every 30 sec.</li>
	<li>Terminate the digestion by centrifugation of the sample for 15 sec at 5,000 x g at 4 &deg;C to remove the pepsin.</li>
	<li>Rapidly freeze the samples in liquid N2 and store at -80 &deg;C for no more than 2 days prior to MALDI-TOF analysis.</li>
	<li>The matrix for MALDI is 5 mg/ml of &alpha;-cyano-4-hydroxycinnamic acid (CHCA) dissolved in a solution (1:1:1) of acetonitrile, ethanol and 0.1 % TFA at pH 2.5 and held at 0 &deg;C.</li>
	<li>Partially thaw each sample quickly and mix 1 &mu;l of each sample with 1 &mu;l of matrix and spot on a 4 &deg;C MALDI target plate.</li>
	<li>Apply a moderate vacuum to the plate to dry the spots in 30 sec prior to MALDI-TOF analysis. The time between thawing of the sample and spectrum retrieval is about 3 min.</li>
	<li>Acquire MALDI-TOF mass spectra with a PerSeptive Biosystems Voyager DE STR. Acquire data at a 2-GHz sampling rate at 100,000 data channels, with a 20,000-V accelerating voltage, and a 65 % grid voltage using delayed extraction with a 180-ns pulse delay. One hundred scans are averaged in 2 min. The MALDI-TOF instrument is not restricted to any brand or any particular software.</li>
	<li>Use the Data Explorer 4.0 computer program to calculate the deuteration of each fragment.</li>
	<li>Calculate the back exchange based on the ratio of the actual incorporated deuteron number at 24 hr of H/D exchange divided by all possible exchangeable sites, which are the backbone amides except the N-terminal amide (Figure 1).</li>
</ol>
4. Representative Results:
The caspase cleavage and autophosphorylation are verified by SDS-PAGE Commassie staining. Inactive Pak2 is a single band of 58 kDa. Caspase cleavage of Pak2 is complete, and produces two fragments, p27 and p34 which migrate separately. The migration of autophosphorylated p27 and p34 shows an overlap on the gel due to the retarded migration of the fully phosphorylated p27 fragment (Figure 2).
H/D exchange experiments are carried out multiple times between 0 and 10 min (Figure 3). As an example, the time course of deuterium incorporation into the m/z peak 1697.85 of inactive Pak2 is composed of multiple isotopic peaks as shown by the shift of the mass envelope over time.
<img alt="Figure 1" src="/files/ftp_upload/3602/3602fig1.jpg" />
Figure 1. Equation for the calculation of deuteration.
<img alt="Figure 2" src="/files/ftp_upload/3602/3602fig2.jpg" />
Figure 2. Autophosphorylation and caspase cleavage of Pak2. Caspase-cleaved Pak2 (left lane), intact inactive Pak2 (middle lane) and caspase-cleaved autophosphorylated Pak2 (right lane) were analyzed by SDS-PAGE and Coommassie blue staining. Adapted from Hsu, YH et al2.
<img alt="Figure 3" src="/files/ftp_upload/3602/3602fig3.jpg" />
Figure 3. Mass spectrum of a time course of deuterium incorporation for inactive Pak2. Inactive Pak2 was subjected to H/D exchange for 0-10 min and analyzed by MALDI-TOF. An example of the expanded isotopic distribution of MALDI-TOF of peak m/z 1697.85 during the time course of H/D exchange is shown. The red dotted line indicates the average of the mass envelope and the shift of the mass envelope shows the deuteration of a peptide.

<ol>
	<li>Anand, G. S., Hughes, C. A., Jones, J. M., Taylor, S. S., Komives, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amide+H/2H+exchange+reveals+communication+between+the+cAMP+and+catalytic+subunit-binding+sites+in+the+R(I)alpha+subunit+of+protein+kinase.">Amide H/2H exchange reveals communication between the cAMP and catalytic subunit-binding sites in the R(I)alpha subunit of protein kinase.</a> A. J. Mol. Biol. 323, 377-386 (2002).</li><li>Hsu, Y. H., Johnson, D. A., Traugh, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+conformational+changes+during+activation+of+protein+kinase+Pak2+by+amide+hydrogen/deuterium+exchange.">Analysis of conformational changes during activation of protein kinase Pak2 by amide hydrogen/deuterium exchange.</a> J. Biol. Chem. 283, 36397-36405 (2008).</li><li>Englander, S. W., Kallenbach, N. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrogen+exchange+and+structural+dynamics+of+proteins+and+nucleic+acids.">Hydrogen exchange and structural dynamics of proteins and nucleic acids.</a> Q. Rev. Biophys. 16, 521-655 (1983).</li><li>Katta, V., Chait, B. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conformational+changes+in+proteins+probed+by+hydrogen-exchange+electrospray-ionization+mass+spectrometry.">Conformational changes in proteins probed by hydrogen-exchange electrospray-ionization mass spectrometry.</a> Rapid Commun. Mass. Spectrom. 5, 214-217 (1991).</li><li>Dennis, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localizing+the+membrane+binding+region+of+Group+VIA+Ca2+-independent+phospholipase+A2+using+peptide+amide+hydrogen/deuterium+exchange+mass+spectrometry.">Localizing the membrane binding region of Group VIA Ca2+-independent phospholipase A2 using peptide amide hydrogen/deuterium exchange mass spectrometry.</a> J. Biol. Chem. 284, 23652-23661 (2009).</li><li>Hsu, Y. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+binding+rigidifies+the+C2+domain+and+the+intradomain+interaction+of+GIVA+phospholipase+A2+as+revealed+by+hydrogen/deuterium+exchange+mass+spectrometry.">Calcium binding rigidifies the C2 domain and the intradomain interaction of GIVA phospholipase A2 as revealed by hydrogen/deuterium exchange mass spectrometry.</a> J. Biol. Chem. 283, 9820-9827 (2008).</li><li>Lee, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+hPAK65+by+caspase+cleavage+induces+some+of+the+morphological+and+biochemical+changes+of+apoptosis.">Activation of hPAK65 by caspase cleavage induces some of the morphological and biochemical changes of apoptosis.</a> Proc. Natl. Acad. Sci. U. S. A. 94, 13642-13647 (1997).</li><li>Rudel, T., Bokoch, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+and+morphological+changes+in+apoptotic+cells+regulated+by+caspase-mediated+activation+of+PAK2.">Membrane and morphological changes in apoptotic cells regulated by caspase-mediated activation of PAK2.</a> Science. 276, 1571-1574 (1997).</li><li>Roig, J., Traugh, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytostatic+p21+G+protein-activated+protein+kinase+gamma-PAK.">Cytostatic p21 G protein-activated protein kinase gamma-PAK.</a> Vitam. Horm. 62, 167-198 (2001).</li><li>Walter, B. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cleavage+and+activation+of+p21-activated+protein+kinase+gamma-PAK+by+CPP32+(caspase+3).+Effects+of+autophosphorylation+on+activity.">Cleavage and activation of p21-activated protein kinase gamma-PAK by CPP32 (caspase 3). Effects of autophosphorylation on activity.</a> J. Biol. Chem. 273, 28733-28739 (1998).</li><li>Clauser, K. R., Baker, P., Burlingame, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+accurate+mass+measurement+(+/-+10+ppm)+in+protein+identification+strategies+employing+MS+or+MS/MS+and+database+searching.">Role of accurate mass measurement (+/- 10 ppm) in protein identification strategies employing MS or MS/MS and database searching.</a> Anal. Chem. 71, 2871-2882 (1999).</li></ol>

No conflicts of interest declared.

Identification of the Pepsin-Digested Fragments is a critical step of the H/D exchange experiment. Incorrect identification of the peptide can lead to a wrong conclusion. Pepsin-digested Pak2 is eluted through a reverse phase HPLC C18 column to obtain a clean background. In our experiments, a total of 40 fractions were collected and subjected to MALDI-TOF. Multiple peptides could be identified in each of the fractions. The peptides were subjected to tandem mass spectrometry (Q-TOF MS/MS and oMALDI MS/MS) to identify their sequences. The MS/MS spectra of a peptide should have at least three product ions to confirm the identification of the peptide.
The Data Explorer 4.0 computer program (Applied Biosystems) performs the initial baseline correction and noise filtration. Each spectrum must be calibrated based on two sequenced peaks. We calibrate our spectrum by the theoretical mass of the undeuterated m/z which are 923.45 and 1697.84. The mass accuracy after calibration can reach 10 ppm. Upon deuteration, the mono-isotope may shift to a higher mass. Unitary step increases to these m/z ratios yielded the enhanced masses associated with deuteration. The peak intensities of the mass envelope will not affect the deuteration level. However, the peak intensities of the isotopic peaks in a particular mass envelope are critical for the calculation of the average mass. The first step of the calculation of the incorporated deuteron is subtracting the peptide mass of the non-deuterated sample from the deuterated sample. Three other factors, the D2O dilution, the residual deuterons in the side chains and the back exchange, will need to be further considered in the calculation (Figure 1). The D2O dilution is the dilution factor of D2O after mixing the sample and D2O buffer to initiate the H/D exchange. The residual deuteron (4.5%) in the side chains of the pepsin-digested peptides needs to be subtracted. The back-exchange is the inevitable loss of deuteration in all deuterated and pepsin-digested peptides in the mass measurement process.
The most solvent accessible peptide (m/z = 1105.60) after 24-hour of H/D exchange represents a full deuteration region. The incorporated deuteron number for each of the peptides is the difference between the centroid of the deuterated and non-deuterated Pak2 peptic peptides. The most highly deuterated peptide m/z 1105 was selected to represent full deuteration when Pak2 was at 24 hr of H/D exchange. The Back Exchange Ratio was calculated by the actual incorporated deuteron number at 24 hr of H/D exchange divided by all possible exchangeable sites at peptide m/z 1105.

<table border="1">
</table>
<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<tr>
			<td>D2O</td>
			<td>Aldrich</td>
			<td>7789-20-0</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>&alpha;-cyano-4-hydroxycinnamic acid</td>
			<td>Sigma</td>
			<td>28166-41-8</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>MALDI-TOF</td>
			<td>PE Biosystems</td>
			<td>Voyager DE STR</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Q-TOF mass spectrometer</td>
			<td>Q-TOF Ultima-Global</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>QSTAR XL oMALDI MS/MS</td>
			<td>Applied Biosystems</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Data Explorer 4.0 computer program</td>
			<td>Applied Biosystems</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

amide hydrogen/deuterium exchange &amp; maldi-tof mass spectrometry analysis of pak2 activation

Amide hydrogen/deuterium exchange (H/D exchange) coupled with mass spectrometry has been widely used to analyze the interface of protein-protein interactions, protein conformational changes, protein dynamics and protein-ligand interactions. H/D exchange on the backbone amide positions has been utilized to measure the deuteration rates of the micro-regions in a protein by mass spectrometry1,2,3. The resolution of this method depends on pepsin digestion of the deuterated protein of interest into peptides that normally range from 3-20 residues. Although the resolution of H/D exchange measured by mass spectrometry is lower than the single residue resolution measured by the Heteronuclear Single Quantum Coherence (HSQC) method of NMR, the mass spectrometry measurement in H/D exchange is not restricted by the size of the protein4. H/D exchange is carried out in an aqueous solution which maintains protein conformation. We provide a method that utilizes the MALDI-TOF for detection2, instead of a HPLC/ESI (electrospray ionization)-MS system5,6. The MALDI-TOF provides accurate mass intensity data for the peptides of the digested protein, in this case protein kinase Pak2 (also called &gamma;-Pak). Proteolysis of Pak 2 is carried out in an offline pepsin digestion. This alternative method, when the user does not have access to a HPLC and pepsin column connected to mass spectrometry, or when the pepsin column on HPLC does not result in an optimal digestion map, for example, the heavily disulfide-bonded secreted Phospholipase A2 (sPLA2). Utilizing this method, we successfully monitored changes in the deuteration level during activation of Pak2 by caspase 3 cleavage and autophosphorylation7,8,9.

Watch this Scientific Journal Video about Amide Hydrogen/Deuterium Exchange & MALDI-TOF Mass Spectrometry Analysis of Pak2 Activation at JoVE.com

Amide Hydrogen/Deuterium Exchange &amp; MALDI-TOF Mass Spectrometry Analysis of Pak2 Activation

MALDI-TOF mass spectrometry was successfully utilized to monitor the amide hydrogen/deuterium exchange in protein kinase Pak2 activation.

Amide hydrogen/deuterium exchange (H/D exchange) coupled with mass spectrometry has been widely used to analyze the interface of protein-protein ...

amide-hydrogendeuterium-exchange-maldi-tof-mass-spectrometry-analysis

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Biology

MALDI Sample Preparation: the Ultra Thin Layer Method

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS) 1,2. The ultra-thin layer method involves the production of a substrate layer of matrix crystals (alpha-cyano-4-hydroxycinnamic acid) on the sample plate, which serves as a seeding ground for subsequent crystallization of a matrix/analyte mixture. Advantages of the ultra-thin layer method over other sample deposition approaches (e.g. dried droplet) are that it provides (i) greater tolerance to impurities such as salts and detergents, (ii) better resolution, and (iii) higher spatial uniformity. This method is especially useful for the accurate mass determination of proteins. The protocol was initially developed and optimized for the analysis of membrane proteins and used to successfully analyze ion channels, metabolite transporters, and receptors, containing between 2 and 12 transmembrane domains 2. Since the original publication, it has also shown to be equally useful for the analysis of soluble proteins. Indeed, we have used it for a large number of proteins having a wide range of properties, including those with molecular masses as high as 380 kDa 3. It is currently our method of choice for the molecular mass analysis of all proteins. The described procedure consistently produces high-quality spectra, and it is sensitive, robust, and easy to implement.

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS).

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption ...

Analyzing Large Protein Complexes by Structural Mass Spectrometry

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an array of dynamic, multi-protein complexes1. To gain a mechanistic understanding of the various cellular processes, it is crucial to determine the structure of such protein complexes, and reveal how their structural organization dictates their function. Many aspects of multi-protein complexes are, however, difficult to characterize, due to their heterogeneous nature, asymmetric structure, and dynamics. Therefore, new approaches are required for the study of the tertiary levels of protein organization.
One of the emerging structural biology tools for analyzing macromolecular complexes is mass spectrometry (MS)2-5. This method yields information on the complex protein composition, subunit stoichiometry, and structural topology. The power of MS derives from its high sensitivity and, as a consequence, low sample requirement, which enables examination of protein complexes expressed at endogenous levels. Another advantage is the speed of analysis, which allows monitoring of reactions in real time. Moreover, the technique can simultaneously measure the characteristics of separate populations co-existing in a mixture. 
Here, we describe a detailed protocol for the application of structural MS to the analysis of large protein assemblies. The procedure begins with the preparation of gold-coated capillaries for nanoflow electrospray ionization (nESI). It then continues with sample preparation, emphasizing the buffer conditions which should be compatible with nESI on the one hand, and enable to maintain complexes intact on the other. We then explain, step-by-step, how to optimize the experimental conditions for high mass measurements and acquire MS and tandem MS spectra. Finally, we chart the data processing and analyses that follow. Rather than attempting to characterize every aspect of protein assemblies, this protocol introduces basic MS procedures, enabling the performance of MS and MS/MS experiments on non-covalent complexes. Overall, our goal is to provide researchers unacquainted with the field of structural MS, with knowledge of the principal experimental tools.

Mass spectrometry has proven to be a valuable tool for analyzing large protein complexes. This method enables insights into the composition, stoichiometry and overall architecture of multi-subunit assemblies. Here, we describe, step-by-step, how to perform a structural mass spectrometry analysis, and characterize macromolecular structures.

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an ...

T-wave Ion Mobility-mass Spectrometry: Basic Experimental Procedures for Protein Complex Analysis

Ion mobility (IM) is a method that measures the time taken for an ion to travel through a pressurized cell under the influence of a weak electric field. The speed by which the ions traverse the drift region depends on their size: large ions will experience a greater number of collisions with the background inert gas (usually N2) and thus travel more slowly through the IM device than those ions that comprise a smaller cross-section. In general, the time it takes for the ions to migrate though the dense gas phase separates them, according to their collision cross-section (&Omega;). 
Recently, IM spectrometry was coupled with mass spectrometry and a traveling-wave (T-wave) Synapt ion mobility mass spectrometer (IM-MS) was released. Integrating mass spectrometry with ion mobility enables an extra dimension of sample separation and definition, yielding a three-dimensional spectrum (mass to charge, intensity, and drift time). This separation technique allows the spectral overlap to decrease, and enables resolution of heterogeneous complexes with very similar mass, or mass-to-charge ratios, but different drift times. Moreover, the drift time measurements provide an important layer of structural information, as &Omega; is related to the overall shape and topology of the ion. The correlation between the measured drift time values and &Omega; is calculated using a calibration curve generated from calibrant proteins with defined cross-sections1.
The power of the IM-MS approach lies in its ability to define the subunit packing and overall shape of protein assemblies at micromolar concentrations, and near-physiological conditions1. Several recent IM studies of both individual proteins2,3 and non-covalent protein complexes4-9, successfully demonstrated that protein quaternary structure is maintained in the gas phase, and highlighted the potential of this approach in the study of protein assemblies of unknown geometry. Here, we provide a detailed description of IMS-MS analysis of protein complexes using the Synapt (Quadrupole-Ion Mobility-Time-of-Flight) HDMS instrument (Waters Ltd; the only commercial IM-MS instrument currently available)10. We describe the basic optimization steps, the calibration of collision cross-sections, and methods for data processing and interpretation. The final step of the protocol discusses methods for calculating theoretical &Omega; values. Overall, the protocol does not attempt to cover every aspect of IM-MS characterization of protein assemblies; rather, its goal is to introduce the practical aspects of the method to new researchers in the field.

Ion mobility-mass spectrometry is an emerging gas-phase technology that separates ions, based on their collision cross-section and mass. The method provides three-dimensional information on the overall topology and shape of protein complexes. Here, we outline a basic procedure for instrument setting and optimization, calibration of drift times, and data interpretation.

Ion mobility (IM) is a method that measures the time taken for an ion to travel through a pressurized cell under the influence of a weak electric field. ...

Direct Analysis of Single Cells by Mass Spectrometry at Atmospheric Pressure

Analysis of biochemicals in single cells is important for understanding cell metabolism, cell cycle, adaptation, disease states, etc. Even the same cell types exhibit heterogeneous biochemical makeup depending on their physiological conditions and interactions with the environment. Conventional methods of mass spectrometry (MS) used for the analysis of biomolecules in single cells rely on extensive sample preparation. Removing the cells from their natural environment and extensive sample processing could lead to changes in the cellular composition. Ambient ionization methods enable the analysis of samples in their native environment and without extensive sample preparation.1 The techniques based on the mid infrared (mid-IR) laser ablation of biological materials at 2.94 &mu;m wavelength utilize the sudden excitation of water that results in phase explosion.2 Ambient ionization techniques based on mid-IR laser radiation, such as laser ablation electrospray ionization (LAESI) and atmospheric pressure infrared matrix-assisted laser desorption ionization (AP IR-MALDI), have successfully demonstrated the ability to directly analyze water-rich tissues and biofluids at atmospheric pressure.3-11 In LAESI the mid-IR laser ablation plume that mostly consists of neutral particulate matter from the sample coalesces with highly charged electrospray droplets to produce ions. Recently, mid-IR ablation of single cells was performed by delivering the mid-IR radiation through an etched fiber. The plume generated from this ablation was postionized by an electrospray enabling the analysis of diverse metabolites in single cells by LAESI-MS.12 This article describes the detailed protocol for single cell analysis using LAESI-MS. The presented video demonstrates the analysis of a single epidermal cell from the skin of an Allium cepa bulb. The schematic of the system is shown in Figure 1. A representative example of single cell ablation and a LAESI mass spectrum from the cell are provided in Figure 2.

Single cell analysis is performed by mass spectrometry on plant and animal cells at atmospheric pressure by using a sharpened optical fiber to sample the cells for laser ablation electrospray ionization (LAESI) mass spectrometry.

Analysis of biochemicals in single cells is important for understanding cell metabolism, cell cycle, adaptation, disease states, etc. Even the same cell ...

Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on Arabidopsis thaliana, a genetically tractable host plant1,2. The accumulation of reactive oxygen species (ROS) in cotyledons inoculated with DC3000 indicates a role of ROS in modulating necrotic cell death during bacterial speck disease of tomato3. Hydrogen peroxide, a component of ROS, is produced after inoculation of tomato plants with Pseudomonas3. Hydrogen peroxide can be detected using a histochemical stain 3'-3' diaminobenzidine (DAB)4. DAB staining reacts with hydrogen peroxide to produce a brown stain on the leaf tissue4. ROS has a regulatory role of the cellular redox environment, which can change the redox status of certain proteins5. Cysteine is an important amino acid sensitive to redox changes. Under mild oxidation, reversible oxidation of cysteine sulfhydryl groups serves as redox sensors and signal transducers that regulate a variety of physiological processes6,7. Tandem mass tag (TMT) reagents enable concurrent identification and multiplexed quantitation of proteins in different samples using tandem mass spectrometry8,9. The cysteine-reactive TMT (cysTMT) reagents enable selective labeling and relative quantitation of cysteine-containing peptides from up to six biological samples. Each isobaric cysTMT tag has the same nominal parent mass and is composed of a sulfhydryl-reactive group, a MS-neutral spacer arm and an MS/MS reporter10. After labeling, the samples were subject to protease digestion. The cysteine-labeled peptides were enriched using a resin containing anti-TMT antibody. During MS/MS analysis, a series of reporter ions (i.e., 126-131 Da) emerge in the low mass region, providing information on relative quantitation. The workflow is effective for reducing sample complexity, improving dynamic range and studying cysteine modifications. Here we present redox proteomic analysis of the Pst DC3000 treated tomato (Rio Grande) leaves using cysTMT technology. This high-throughput method has the potential to be applied to studying other redox-regulated physiological processes.

Reactive oxygen species level is elevated when cells encounter stress conditions. Here we show the example of 3'-3' diaminobenzidine staining as well as cysTMT labeling and mass spectrometry to profile the redox proteome in Pseudomonas syringae treated tomato leaves.

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on ...

Use of MALDI-TOF Mass Spectrometry and a Custom Database to Characterize Bacteria Indigenous to a Unique Cave Environment (Kartchner Caverns, AZ, USA)

MALDI-TOF mass spectrometry has been shown to be a rapid and reliable tool for identification of bacteria at the genus and species, and in some cases, strain levels. Commercially available and open source software tools have been developed to facilitate identification; however, no universal/standardized data analysis pipeline has been described in the literature. Here, we provide a comprehensive and detailed demonstration of bacterial identification procedures using a MALDI-TOF mass spectrometer. Mass spectra were collected from 15 diverse bacteria isolated from Kartchner Caverns, AZ, USA, and identified by 16S rDNA sequencing. Databases were constructed in BioNumerics 7.1. Follow-up analyses of mass spectra were performed, including cluster analyses, peak matching, and statistical analyses. Identification was performed using blind-coded samples randomly selected from these 15 bacteria. Two identification methods are presented: similarity coefficient-based and biomarker-based methods. Results show that both identification methods can identify the bacteria to the species level.

Use of MALDI-TOF Mass Spectrometry and a Custom Database to Characterize Bacteria Indigenous to a Unique Cave Environment Kartchner Caverns, AZ, USA

This work details procedures for rapid identification of bacteria using MALDI-TOF MS. The identification procedures include spectrum acquisition, database construction, and follow up analyses. Two identification methods, similarity coefficient-based and biomarker-based methods, are presented.

MALDI-TOF mass spectrometry has been shown to be a rapid and reliable tool for identification of bacteria at the genus and species, and in some cases, ...

Selected Reaction Monitoring Mass Spectrometry for Absolute Protein Quantification

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This protocol provides step-by-step instructions for the development and application of quantitative assays using selected reaction monitoring (SRM) mass spectrometry (MS). First, likely quantotypic target peptides are identified based on numerous criteria. This includes identifying proteotypic peptides, avoiding sites of posttranslational modification, and analyzing the uniqueness of the target peptide to the target protein. Next, crude external peptide standards are synthesized and used to develop SRM assays, and the resulting assays are used to perform qualitative analyses of the biological samples. Finally, purified, quantified, heavy isotope labeled internal peptide standards are prepared and used to perform isotope dilution series SRM assays. Analysis of all of the resulting MS data is presented. This protocol was used to accurately assay the absolute abundance of proteins of the chemotaxis signaling pathway within RAW 264.7 cells (a mouse monocyte/macrophage cell line). The quantification of Gi2 (a heterotrimeric G-protein &#945;-subunit) is described in detail.

This protocol describes how to perform absolute quantification assays of target proteins within complex biological samples using selected reaction monitoring. It was used to accurately quantify proteins of the mouse macrophage chemotaxis signaling pathway. Target peptide selection, assay development, and qualitative and quantitative assays are described in detail.

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This ...

Enrichment of Extracellular Matrix Proteins from Tissues and Digestion into Peptides for Mass Spectrometry Analysis

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of cell proliferation, survival, migration, etc. The ECM plays important roles in development and in diverse pathologies including cardio-vascular and musculo-skeletal diseases, fibrosis, and cancer. Thus, characterizing the composition of ECMs of normal and diseased tissues could lead to the identification of novel prognostic and diagnostic biomarkers and potential novel therapeutic targets. However, the very nature of ECM proteins (large in size, cross-linked and covalently bound, heavily glycosylated) has rendered biochemical analyses of ECMs challenging. To overcome this challenge, we developed a method to enrich ECMs from fresh or frozen tissues and tumors that takes advantage of the insolubility of ECM proteins. We describe here in detail the decellularization procedure that consists of sequential incubations in buffers of different pH and salt and detergent concentrations and that results in 1) the extraction of intracellular (cytosolic, nuclear, membrane and cytoskeletal) proteins and 2) the enrichment of ECM proteins. We then describe how to deglycosylate and digest ECM-enriched protein preparations into peptides for subsequent analysis by mass spectrometry.

This protocol describes a procedure for enriching ECM proteins from tissues or tumors and deglycosylating and digesting the ECM-enriched preparations into peptides to analyze their protein composition by mass spectrometry.

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of ...

Applications of the Single-probe: Mass Spectrometry Imaging and Single Cell Analysis under Ambient Conditions

Mass spectrometry imaging (MSI) and in-situ single cell mass spectrometry (SCMS) analysis under ambient conditions are two emerging fields with great potential for the detailed mass spectrometry (MS) analysis of biomolecules from biological samples. The single-probe, a miniaturized device with integrated sampling and ionization capabilities, is capable of performing both ambient MSI and in-situ SCMS analysis. For ambient MSI, the single-probe uses surface micro-extraction to continually conduct MS analysis of the sample, and this technique allows the creation of MS images with high spatial resolution (8.5 &#181;m) from biological samples such as mouse brain and kidney sections. Ambient MSI has the advantage that little to no sample preparation is needed before the analysis, which reduces the amount of potential artifacts present in data acquisition and allows a more representative analysis of the sample to be acquired. For in-situ SCMS, the single-probe tip can be directly inserted into live eukaryotic cells such as HeLa cells, due to the small sampling tip size (&lt; 10 &#181;m), and this technique is capable of detecting a wide range of metabolites inside individual cells at near real-time. SCMS enables a greater sensitivity and accuracy of chemical information to be acquired at the single cell level, which could improve our understanding of biological processes at a more fundamental level than previously possible. The single-probe device can be potentially coupled with a variety of mass spectrometers for broad ranges of MSI and SCMS studies.

Here, we present protocols to perform both ambient mass spectrometry imaging (MSI) of tissues and in-situ live single cell MS (SCMS) analysis using the single-probe, which is a miniaturized multifunctional device for MS analysis.

Mass spectrometry imaging (MSI) and in-situ single cell mass spectrometry (SCMS) analysis under ambient conditions are two emerging fields with great ...

Complete Workflow for Analysis of Histone Post-translational Modifications Using Bottom-up Mass Spectrometry: From Histone Extraction to Data Analysis

Nucleosomes are the smallest structural unit of chromatin, composed of 147 base pairs of DNA wrapped around an octamer of histone proteins. Histone function is mediated by extensive post-translational modification by a myriad of nuclear proteins. These modifications are critical for nuclear integrity as they regulate chromatin structure and recruit enzymes involved in gene regulation, DNA repair and chromosome condensation. Even though a large part of the scientific community adopts antibody-based techniques to characterize histone PTM abundance, these approaches are low throughput and biased against hypermodified proteins, as the epitope might be obstructed by nearby modifications. This protocol describes the use of nano liquid chromatography (nLC) and mass spectrometry (MS) for accurate quantification of histone modifications. This method is designed to characterize a large variety of histone PTMs and the relative abundance of several histone variants within single analyses. In this protocol, histones are derivatized with propionic anhydride followed by digestion with trypsin to generate peptides of 5 - 20 aa in length. After digestion, the newly exposed N-termini of the histone peptides are derivatized to improve chromatographic retention during nLC-MS. This method allows for the relative quantification of histone PTMs spanning four orders of magnitude.

This protocol outlines a fully integrated workflow for characterizing histone post-translational modifications using mass spectrometry (MS). The workflow includes histone purification from cell cultures or tissues, histone derivatization and digestion, MS analysis using nano-flow liquid chromatography and instructions for data analysis. The protocol is designed for completion within 2 - 3 days.

Nucleosomes are the smallest structural unit of chromatin, composed of 147 base pairs of DNA wrapped around an octamer of histone proteins. Histone ...