The authors thank the Sharon group members for their critical review, and for their contributions to the manuscript. We are grateful for the support of the Morasha and Bikura Programs, the Israel Science Foundation (Grant Nos. 1823/07 and 378/08), the Josef Cohn Minerva Center for Biomembrane Research, the Chais Family Fellows Program for New Scientists, the Abraham and Sonia Rochlin Foundation; the Wolfson Family Charitable Trust; the Helen and Milton A. Kimmelman Center for Biomolecular Structure and Assembly; the estate of Shlomo and Sabine Beirzwinsky; Meil de Botton Aynsley, and Karen Siem, UK. We are grateful to Dan Tawfik and Itamar Yadid for giving us the lectin variant sample.

Part 1: Preparation of gold-coated capillaries for nanoflow electrospray ionization
Analysis of non-covalent complexes is usually performed by means of nanoflow electrospray ionization (nESI)6, using glass or quartz capillaries which have been pulled to a fine tip (~1 &#956;m inner diameter), and coated with conductive material (usually gold). Such capillaries are available ready-to-use from commercial sources (New Objective or Proxeon); however, it can be more cost-effective to prepare them in-house:
<ol>
	<li>Stick two strips of a double-sided adhesive pad to the bottom of a Petri dish, 2 cm apart. Place a glass rod (8 cm x 5 mm) in the center of one of the pads. The adhesive pad will hold the capillaries in place, and the glass rod will support the prepared capillaries, and keep the tips from breaking (Figure&#160;1).</li>
	<li>Use borosilicate glass capillaries, 1.0 mm OD x 0.78 mm i.d. (we use packs of 500 thin wall borosilicate capillaries from Warner Instruments, cat. no. G100TF- 4). Insert one capillary into the needle puller (we use model P-97 from the Sutter Instrument Co.). Clamp the capillary gently in place, and adjust its position so that it lies in the center of the needle puller&#39;s heating filament. Tighten the clamps gently, until the capillary is held firm at both ends.</li>
	<li>Pull the capillary, using a predefined program. Every capillary pulled will give rise to two final, shaped capillaries. The process of programming the puller is one of trial-and-error, until an acceptable tip shape is obtained. We use the following program:
	<table border="1">
		<tbody>
			<tr>
				<td>Step</td>
				<td>Heat</td>
				<td>Pull</td>
				<td>Vell</td>
				<td>Time</td>
			</tr>
			<tr>
				<td>1</td>
				<td>750</td>
				<td>-</td>
				<td>15</td>
				<td>80</td>
			</tr>
			<tr>
				<td>2</td>
				<td>700</td>
				<td>-</td>
				<td>15</td>
				<td>50</td>
			</tr>
			<tr>
				<td>3</td>
				<td>750</td>
				<td>200</td>
				<td>20</td>
				<td>80</td>
			</tr>
		</tbody>
	</table>
	</li>
	<li>Remove the pulled capillaries from the instrument and inspect the tips, discarding any that are deformed or broken. Use blunt tweezers (we use positioning tweezers from C.K. Precision), to place the capillaries in the Petri dish. The base of the capillary should be attached to the adhesive pad and the upper part should lean on the glass rod, with the tip pointing up.</li>
	<li>Once the Petri dish is full (about 80 capillaries fit into a 10 cm-diameter dish), insert the plate into the gold sputter coater (we use model no. EMS550, from EMS). Make sure that the gas supply is chosen according to the manufacturer&#39;s instructions, and activate a predefined coating cycle (we use Argon pressure 4 psi, at a vacuum pressure of 5 x 10-2 mbar, a current of 45mA, and a coating time of 1 min, for 3-6 cycles, until the capillaries are evenly golden).</li>
</ol>
Part 2: Sample preparation
<ol>
	<li>Low micromolar concentrations of sample are required (1 - 20 &#956;M). If necessary, concentrate the sample using centrifugal ultrafiltration devices (e.g., Vivaspin from Sartorius, or NanoSep from Pall Corporation). It is recommended that you verify the adsorption of the protein complex to the device membrane, before use.</li>
	<li>Often, the purification buffers or storage solutions of the protein complex are not compatible with nESI, for which only volatile solutions may be used. Therefore, buffer exchange is necessary. This critical step, in which all traces of salts, buffer molecules or any other non-volatile adducts such as glycerol, DTT, or EDTA are removed, determines the quality of the spectra. Usually, aqueous ammonium acetate solution is used, at a concentration of between 5 mM and 1 M, and at 6-8 pH. Buffer exchange can be performed using a microcentrifuge gel filtration column (e.g., Micro Bio-Spin 6 chromatography columns from Bio-Rad). This step can be repeated 1-3 times with minimal dilution (less than a factor of 1.3 per device5), until maximal exchange is achieved. If both concentration and buffer exchange are required, these may be done together, using centrifugal ultrafiltration (e,g., Vivaspin from Sartorius, or NanoSep from Pall Corporation).</li>
</ol>
Part 3: Calibrating the mass spectrometer for high mass measurements
Most of the experiments conducted on multi-protein complexes are performed using a nano electrospray quadrupole-time-of-flight (Q-ToF) instrument. It is suggested that you use a quadrupole mass filter adjusted to low frequencies, to enable transmission and mass analysis of ions with high m/z values7,8. It is also recommended that gas inlets7,8 or sleeves9 be added to the instrument in the first ion guide, to enable pressure control at the first vacuum stage. The latter enables optimization of the transmission, and desolvation of very large ions7-9. Currently, commercial ESI-ToF and Q-ToF instruments are available from several manufacturers (for example, Waters, SCIEX, Bruker, or Agilent) which can be modified relatively easily and cost-effectively, for native MS applications7,8. It is possible, however, to use standard ToF or QToF configurations on instruments such as LCT or QToF1 (Waters) to acquire mass spectra for complexes up to 1 MDa, without the need for hardware modifications5.
The protocol outlined below was conducted on a Synapt instrument (Waters).
<ol>
	<li>Prepare 100 mg/ml solution of CsI in purified water. CsI is used for high mass calibration as the singly charged salt clusters, (CsI)nCs+ extend over a wide mass range, from 393 m/z to well over 10,000 (Figure&#160;2).</li>
	<li>Using blunt tweezers, take a coated capillary from the Petri dish and load 2&#956;L of the CsI solution into the capillary, using an Eppendorf GeLoader tip.</li>
	<li>Insert the capillary into the capillary holder, and adjust the capillary in such a way that the tip is approximately 10 mm away from the edge of the holder.</li>
	<li>Slide the solution towards the tip of the capillary either manually, or using a spin down adaptor.</li>
	<li>Place the capillary under an optical microscope and trim the tip, using the sharp AA-type tweezers.</li>
	<li>Connect the capillary holder to the nanoflow ES interface. Rotate the x-y-z stage backwards to avoid damaging the capillary, and push the stage into its activated position. The capillary should be placed 1 - 10 mm from the cone orifice.</li>
	<li>Apply capillary voltage (1,050-1,400 V) and low nanoflow pressure (0.00-0.03 bar) until spray is initiated, then try to reduce the nanoflow pressure to a minimal value.</li>
	<li>Optimize the signal intensity by adjusting the location of the x-y-z stage, the capillary voltage, the nanoflow pressure, and the desolvation gas flow.</li>
	<li>To detect a wide mass range of CsI peak series, the accelerating voltages should be optimized (we use the following parameters: capillary 1.3-1.7 kV, sample cone 80-150 V, extraction cone 1-3 V).</li>
	<li>To optimize the transmission of high mass ions, which require gentle desolvation conditions, the backing pressure in the initial vacuum stage, between the source and the analyzer, should be raised. This can be achieved by carefully reducing the conductance of the source vacuum line to the scroll pump, by partially closing the isolation valve (SpeediValve). In order to define the optimal point, the latter should be done while monitoring the effect on signal intensity (we usually use 3.0-6.5 mBar).</li>
	<li>Collect about 30 scans at 1 scan/sec, at an m/z range of between 1,000 to 15,000 m/z.</li>
	<li>After acquisition, calibrate the TOF using the appropriate calibration table.</li>
</ol>
Part 4: MS analysis of intact protein complexes
<ol>
	<li>Load your sample, as described (Part 3, Sections 2-5), and initiate spray.</li>
	<li>Start by initial optimization of the spray (Part 3, Sections 6-9), until the signal is detected. The exact position of the charge states is protein-dependent; however, it can be predicted, using the relationship between ion mass and average charge state, thus: (Zav)10: Zav = 0.0778&#8730;(m) , in which m is the mass of the complex in Daltons. To prevent complex dissociation, do not heat the ion source: either switch the heater off, or keep the temperature below 40 &#176;C.</li>
	<li>Vary the capillary position, sample cone and extractor voltages to maximize ion transmission, and check the resulting change in the spectra. A possible starting point could be cone voltage 100 V; extractor cone: 1 V; capillary voltage 1.5 kV. Optimize these parameters, in combination with the nanoflow pressure.</li>
	<li>To improve desolvation and strip off residual water and buffer components, increase the bias voltage and the gas pressure in the collision cell. This step should be performed carefully, to prevent dissociation of the complex. Typical bias voltages are within the range of 10-100 V, with a trap gas flow of 1-10 ml/min (Figure&#160;3). Manual adjustment of the RF setting and quadrupole profile may improve the transmission of high mass ions.</li>
	<li>Adjust Trap and Transfer collision energies. Often, higher voltages are required for transmission of high mass ions (usually within the range of 10-30 V). At this point, it is important to avoid collision-induced dissociation of the complex.</li>
	<li>After a stable signal is reached, it is recommended that the nanoflow pressure and capillary voltage be decreased to minimal values, while maintaining stable spray.</li>
</ol>
Part 5: Tandem mass spectrometry: dissociating protein complexes
<ol>
	<li>Once an optimum and stable signal has been obtained, select a precursor ion. Set the mass center and isolation width (we generally use a LM resolution of 12, and a HM resolution of between 13 and 15). Use a wide mass range to detect the high mass/low charge dissociation products. It is recommended that you set the m/z range to the maximum level, and then reduce it to the desired values. We further suggest superimposing the MS and MS/MS spectra, to validate the isolation of the chosen precursor ion.</li>
	<li>To perform MS/MS, dissociate the precursor ion by increasing the collision energy (CE) and the pressure on the collision cell. Increase either Trap or Transfer CE gradually, in steps of 10-20 V, and elevate the collision gas pressure to 0-5 ml/min (common values). Monitor changes in the spectra until optimal activation conditions are reached. High activation energy may induce the dissociation of one or more subunits from the intact complex, and clarify the interaction affinities of different subunits. We typically use Argon as a collision gas in the Trap/Transfer cells, although benefits of using a heavier gas (e.g., Xe or SF6) have been reported; however, these gases are substantially more expensive11 (Figure&#160;3).</li>
	<li>It is recommended that more than one charge state be selected for MS/MS analysis. In the case of overlapping components, acquiring a set of tandem MS spectra will assist in resolving the charge series of the different populations. Moreover, higher charge states will dissociate more easily, compared to lower charge states12.</li>
	<li>In addition to the characterization of the intact complex, it is suggested that smaller subcomplexes in solution will be generated, under mild denaturing conditions. The MS and MS/MS analyses of subcomplexes form the basis for defining the subunit architecture of the complex13. For partial disruption of subunit-subunit interactions, gradually add organic solvents (e.g. methanol, isopropanol, or acetonitrile) up to a concentration of 50%, or change the pH of the solution by adding ammonia or formic acid (up to a concentration of 4%).</li>
	<li>To determine the masses of the individual subunits that compose the complex, it is important to acquire a spectrum under denaturing conditions. This can be carried out using Zip-Tip C4 (Millipore) with a 25:75 water/acetonitrile ratio, with 1% formic acid as the elution solvent.</li>
</ol>
Part 6: Data processing and analysis
<ol>
	<li>Data is analyzed offline, using a spectral analysis programs for MS spectra. We normally use the MassLynx program (Waters).</li>
	<li>For spectra that span a wide m/z range, we recommend that different schemes of smoothing and centroid parameters be applied for the high and low m/z regions, to reflect the difference in peak resolution of these regions.</li>
	<li>To identify charge series and calculate charge states and masses, the &#34;manual find&#34; function of MassLynx can be applied. In cases of complex mass spectra with overlapping components, manual calculation of masses may be easier.</li>
	<li>To facilitate these assignment processes, additional software may be used; for example: MaxEnt for spectra deconvolution14, SOMMS for peak fitting and simulation15, and SUMMIT for assigning the composition and stoichiometry of protein subcomplexes and generating protein interaction networks13,16,17.</li>
</ol>
Part 7: Representative Results
<img alt="Figure 1" src="/files/ftp_upload/1954/1954fig1.jpg" /> 
Figure 1. Preparing gold-coated nano-electrospray capillaries. 
A. Attach two double-sided adhesive strips to a Petri dish, 2 cm apart. To support the prepared capillaries, place a glass rod (8 cm x 5 mm) in the center of one of the pads. B. Stick the blunt end of the prepared capillaries to the adhesive pad, and lean the tip on the glass rod. C. Once the Petri dish is filled with the prepared capillaries, coat them with gold until a thin film of gold is evenly deposited on the external surface of the capillaries.
<img alt="Figure 2" src="/files/ftp_upload/1954/1954fig2.jpg" /> 
Figure 2. High mass calibration using cesium iodide ions. 
The large and monisotopic clusters of CsI have made it the compound of choice for calibrating mass spectrometers for high mass analysis. The series of equally spaced peaks extend over a wide range, from m/z 393 to well over 10,000. They are assigned to singly charged salt clusters of the general composition (CsI)nCs+. Additional signals between the major peaks are caused by double- and triple-charged species of the series; [(CsI)nCs2]2+ and [(CsI)nCs3]3+, respectively. Increasing the pressure at the initial vacuum stage is essential for detecting the high mass clusters. The effect of pressure on the high mass peaks is demonstrated in Panels A. and B. with pressure readbacks of 1.2 and 5.3, respectively. C. Expansion of the mass spectrum shown at B.
<img alt="Figure 3" src="/files/ftp_upload/1954/1954fig3.jpg" /> 
Figure 3. Nanoflow electrospray mass spectra of a pentameric lectin. 
A. Mass spectrometry of a lectin variant complex (derived from Lib1-B7 by directed evolution18) gives rise to a charge state distributions between 3,000 and 5,000 m/z; however, due to inadequate desolvation of the ions, the peaks are broad. Comparison of panel A. and B. show the effect of increasing the bias voltage from 4 V (A.) to 15 V (B.) on the peak width. This increase in accelerating conditions causes the stripping of residual water and buffer components, yielding a highly resolved spectrum. The measured mass (60,240 &#177; 38 Da) corresponds to a pentameric complex. C. The +15 charge state was then selected for tandem MS analysis (shaded in grey in Panel B.) D. The increase in collision energy causes the release of a highly charged monomer, centered at 1,664 m/z, and a stripped tetrameric complex, in the range of 5,000&#8211;8,000 m/z. All spectra were obtained from a sample containing 20 &#956;M of solution in 0.5 M ammonium acetate.

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No conflicts of interest declared.

To acquire high quality spectra attention should be given to the sample preparation steps, which include sample concentration and buffer exchange. Over diluted samples will yield a low signal, whereas highly concentrated samples can be rather viscous, and block the electrospray needle. Moreover, solution additives such as salts, glycerol, detergents, metal ions, and reducing agents (DTT or &#946;-mercaptoethanol), tend to adhere to the outer surface of the proteins, and cause broadening of the peaks. Therefore, in order to achieve well-resolved peaks, these components should be added at the lowest concentrations possible.
Another key parameter is the position of the nanoflow capillary, relative to the mass spectrometer orifice. Finding the &#34;sweet spot&#34; might be challenging for inexperienced users; nevertheless, it significantly influences the quality of the spectra. It is also important to examine the nanoflow capillary prior to initiation of spray. The droplet size is a function of the tip diameter, which should be ~1 &#956;m. Small droplets lead to more efficient ionization, and would therefore be advantageous. It is also important to validate that there are no air bubbles within the capillary that could block the flow, and that the gold coating is not stripped from the capillary during acquisition; if so, further trim the tip. Keep in mind that an excessive amount of sample will increase the possibility of optimizing MS conditions such as the capillary voltage, accelerating voltages, pressure, and collision energy.
Overall, the procedures explained in the protocol have been used to determine the composition, stoichiometry and architecture of numerous protein complexes (see reviews 2,3,4). Analysis of large MDa complexes such as the ribosome19 and highly ordered virus capsids20-22, in defining substrate binding to molecular machines23-25, or the characterization of subunit interaction networks26,17,16,17,27,28 serve as but a few examples of the value of this approach.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
			<div>
			Sample requirements:	
<table border="1">
 <tr>
 <td>Sample</td>
 <td>Requirement </td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Volume</td>
 <td>1-2 &mu;L</td>
 <td>Per capillary</td>
 </tr>
 <tr>
 <td>Concentration</td>
 <td>1-20&mu;M</td>
 <td>Per complex</td>
 </tr>
 <tr>
 <td>Buffer</td>
 <td>Aquanos volatile buffer such as ammonium acetate at pH= 6- 8</td>
 <td>Typically 5 mM-1M</td>
 </tr>
 <tr>
 <td>Detergent</td>
 <td>Minimal </td>
 <td>Clusters of detergent molecules produce broad and unresolved peaks </td>
 </tr>
 <tr>
 <td>Glycerol</td>
 <td>Minimal (up to 5%)</td>
 <td>Adheres nonspecifically to proteins and, consequently, broad peaks are observed</td>
 </tr>
 <tr>
 <td>Organic solvents</td>
 <td>Up to 50%</td>
 <td>Might denature proteins complexes </td>
 </tr>
 <tr>
 <td>Acids</td>
 <td>Up to 4%</td>
 <td>Denature protein complexes </td>
 </tr>
 <tr>
 <td>Salts</td>
 <td>Minimal</td>
 <td>Salt adducts lead to broad and unresolved peaks</td>
 </tr>
 <tr>
 <td>DTT</td>
 <td>Minimal</td>
 <td>1-2 &mu;M can be present</td>
 </tr>
 <tr>
 <td>Chelating agents</td>
 <td>Minimal</td>
 <td>Above 250 &mu;M lead to extensive adduct formation</td>
 </tr>
</table>		</div>

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.

Watch this Scientific Journal Video about Analyzing Large Protein Complexes by Structural Mass Spectrometry at JoVE.com

Analyzing Large Protein Complexes by Structural Mass Spectrometry

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 ...

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24.0K Views.  Weizmann Institute of Science. 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.

Video: Analyzing Large Protein Complexes by Structural Mass Spectrometry

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 ...

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. ...

Atmospheric-pressure Molecular Imaging of Biological Tissues and Biofilms by LAESI Mass Spectrometry

Ambient ionization methods in mass spectrometry allow analytical investigations to be performed directly on a tissue or biofilm under native-like experimental conditions. Laser ablation electrospray ionization (LAESI) is one such development and is particularly well-suited for the investigation of water-containing specimens. LAESI utilizes a mid-infrared laser beam (2.94 &mu;m wavelength) to excite the water molecules of the sample. When the ablation fluence threshold is exceeded, the sample material is expelled in the form of particulate matter and these projectiles travel to tens of millimeters above the sample surface. In LAESI, this ablation plume is intercepted by highly charged droplets to capture a fraction of the ejected sample material and convert its chemical constituents into gas-phase ions. A mass spectrometer equipped with an atmospheric-pressure ion source interface is employed to analyze and record the composition of the released ions originating from the probed area (pixel) of the sample. A systematic interrogation over an array of pixels opens a way for molecular imaging in the microprobe analysis mode. A unique aspect of LAESI mass spectrometric imaging is depth profiling that, in combination with lateral imaging, enables three-dimensional (3D) molecular imaging. With current lateral and depth resolutions of ~100 &mu;m and ~40 &mu;m, respectively, LAESI mass spectrometric imaging helps to explore the molecular structure of biological tissues. Herein, we review the major elements of a LAESI system and provide guidelines for a successful imaging experiment.

Laser ablation electrospray ionization (LAESI) is an atmospheric-pressure ion source for mass spectrometry. In the imaging mode, a mid-infrared laser probes the distributions of molecules across a tissue section or a biofilm. This technique presents a new approach for diverse bioanalytical studies carried out under native experimental conditions.

Ambient ionization methods in mass spectrometry allow analytical investigations to be performed directly on a tissue or biofilm under native-like ...

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 of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry (CCMS)

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity chromatography 1 or Activity Based Protein Profiling 2. Trifunctional Capture Compounds (CCs, Figure 1A) 3 are the basis for a generic approach, in which the initial equilibrium-driven interaction between a small molecule probe (the selectivity function, here S-adenosyl-L-homocysteine, SAH, Figure 1A) and target proteins is irreversibly fixed upon photo-crosslinking between an independent photo-activable reactivity function (here a phenylazide) of the CC and the surface of the target proteins. The sorting function (here biotin) serves to isolate the CC - protein conjugates from complex biological mixtures with the help of a solid phase (here streptavidin magnetic beads). Two configurations of the experiments are possible: "off-bead" 4 or the presently described "on-bead" configuration (Figure 1B). The selectivity function may be virtually any small molecule of interest (substrates, inhibitors, drug molecules).
S-Adenosyl-L-methionine (SAM, Figure 1A) is probably, second to ATP, the most widely used cofactor in nature 5, 6. It is used as the major methyl group donor in all living organisms with the chemical reaction being catalyzed by SAM-dependent methyltransferases (MTases), which methylate DNA 7, RNA 8, proteins 9, or small molecules 10. Given the crucial role of methylation reactions in diverse physiological scenarios (gene regulation, epigenetics, metabolism), the profiling of MTases can be expected to become of similar importance in functional proteomics as the profiling of kinases. Analytical tools for their profiling, however, have not been available. We recently introduced a CC with SAH as selectivity group to fill this technological gap (Figure 1A).
SAH, the product of SAM after methyl transfer, is a known general MTase product inhibitor 11. For this reason and because the natural cofactor SAM is used by further enzymes transferring other parts of the cofactor or initiating radical reactions as well as because of its chemical instability 12, SAH is an ideal selectivity function for a CC to target MTases. Here, we report the utility of the SAH-CC and CCMS by profiling MTases and other SAH-binding proteins from the strain DH5&alpha; of Escherichia coli (E. coli), one of the best-characterized prokaryotes, which has served as the preferred model organism in countless biochemical, biological, and biotechnological studies. Photo-activated crosslinking enhances yield and sensitivity of the experiment, and the specificity can be readily tested for in competition experiments using an excess of free SAH.

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry CCMS

Capture Compounds are trifunctional small molecules to reduce the complexity of the proteome by functional reversible small molecule-protein interaction followed by photo-crosslinking and purification. Here we use a Capture Compound with S-adenosyl-L-homocysteine-binding as selectivity function to isolate methyltransferases from an Escherichia coli whole cell lysate and identify them by MS.

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity ...

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 ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in current research and a plethora of methods for their investigation is available1. Protein-protein interactions often are highly dynamic and may depend on subcellular localization, post-translational modifications and the local protein environment2. Therefore, they should be investigated in their natural environment, for which co-immunoprecipitation approaches are the method of choice3. Co-precipitated interaction partners are identified either by immunoblotting in a targeted approach, or by mass spectrometry (LC-MS/MS) in an untargeted way. The latter strategy often is adversely affected by a large number of false positive discoveries, mainly derived from the high sensitivity of modern mass spectrometers that confidently detect traces of unspecifically precipitating proteins. A recent approach to overcome this problem is based on the idea that reduced amounts of specific interaction partners will co-precipitate with a given target protein whose cellular concentration is reduced by RNAi, while the amounts of unspecifically precipitating proteins should be unaffected. This approach, termed QUICK for QUantitative Immunoprecipitation Combined with Knockdown4, employs Stable Isotope Labeling of Amino acids in Cell culture (SILAC)5 and MS to quantify the amounts of proteins immunoprecipitated from wild-type and knock-down strains. Proteins found in a 1:1 ratio can be considered as contaminants, those enriched in precipitates from the wild type as specific interaction partners of the target protein. Although innovative, QUICK bears some limitations: first, SILAC is cost-intensive and limited to organisms that ideally are auxotrophic for arginine and/or lysine. Moreover, when heavy arginine is fed, arginine-to-proline interconversion results in additional mass shifts for each proline in a peptide and slightly dilutes heavy with light arginine, which makes quantification more tedious and less accurate5,6. Second, QUICK requires that antibodies are titrated such that they do not become saturated with target protein in extracts from knock-down mutants.
Here we introduce a modified QUICK protocol which overcomes the abovementioned limitations of QUICK by replacing SILAC for 15N metabolic labeling and by replacing RNAi-mediated knock-down for affinity modulation of protein-protein interactions. We demonstrate the applicability of this protocol using the unicellular green alga Chlamydomonas reinhardtii as model organism and the chloroplast HSP70B chaperone as target protein7 (Figure 1). HSP70s are known to interact with specific co-chaperones and substrates only in the ADP state8. We exploit this property as a means to verify the specific interaction of HSP70B with its nucleotide exchange factor CGE19.

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

We present a variation of the QUICK (QUantitative Immunoprecipitation Combined with Knockdown) approach that was introduced previously to distinguish between true and false protein-protein interactions. Our approach is based on 15N metabolic labeling, the modulation of affinities of protein-protein interactions by the presence/absence of ATP, immunoprecipitation, and quantitative mass spectrometry.

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in ...

Protein Purification-free Method of Binding Affinity Determination by Microscale Thermophoresis

Quantitative characterization of protein interactions is essential in practically any field of life sciences, particularly drug discovery. Most of currently available methods of KD determination require access to purified protein of interest, generation of which can be time-consuming and expensive. We have developed a protocol that allows for determination of binding affinity by microscale thermophoresis (MST) without purification of the target protein from cell lysates. The method involves overexpression of the GFP-fused protein and cell lysis in non-denaturing conditions. Application of the method to STAT3-GFP transiently expressed in HEK293 cells allowed to determine for the first time the affinity of the well-studied transcription factor to oligonucleotides with different sequences. The protocol is straightforward and can have a variety of application for studying interactions of proteins with small molecules, peptides, DNA, RNA, and proteins.

Microscale thermophoresis (MST) can be widely used for determination of binding affinity without purification of the target protein from cell lysates. The protocol involves overexpression of the GFP-fused protein, cell lysis in non-denaturing conditions, and detection of MST signal in the presence of varying concentrations of the ligand.

Quantitative characterization of protein  interactions is essential in practically any field of life sciences,  particularly drug discovery. Most of ...

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The introduced protocol provides a tool for the analysis of multiprotein complexes in the thylakoid membrane, by revealing insights into complex composition under different conditions. In this protocol the approach is demonstrated by comparing the composition of the protein complex responsible for cyclic electron flow (CEF) in Chlamydomonas reinhardtii, isolated from genetically different strains. The procedure comprises the isolation of thylakoid membranes, followed by their separation into multiprotein complexes by sucrose density gradient centrifugation, SDS-PAGE, immunodetection and comparative, quantitative mass spectrometry (MS) based on differential metabolic labeling (14N/15N) of the analyzed strains. Detergent solubilized thylakoid membranes are loaded on sucrose density gradients at equal chlorophyll concentration. After ultracentrifugation, the gradients are separated into fractions, which are analyzed by mass-spectrometry based on equal volume. This approach allows the investigation of the composition within the gradient fractions and moreover to analyze the migration behavior of different proteins, especially focusing on ANR1, CAS, and PGRL1. Furthermore, this method is demonstrated by confirming the results with immunoblotting and additionally by supporting the findings from previous studies (the identification and PSI-dependent migration of proteins that were previously described to be part of the CEF-supercomplex such as PGRL1, FNR, and cyt f). Notably, this approach is applicable to address a broad range of questions for which this protocol can be adopted and e.g. used for comparative analyses of multiprotein complex composition isolated from distinct environmental conditions.

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The described comparative, quantitative proteomic approach aims at obtaining insights into the composition of multiprotein complexes&#160;under different conditions and is demonstrated by comparing genetically different strains. For quantitative analysis equal volumes of different fractions from a sucrose density gradient are mixed and analyzed by mass spectrometry.