M.N-S acknowledges support from the National Science Foundation, RUI grant CHE 1011859. The authors gratefully acknowledge the Triad Mass Spectrometry Facility at the University of North Carolina at Greensboro for use of the Thermo Fisher Scientific LTQ Orbitrap XL mass spectrometer. The authors thank Daniel Todd, Vincent Sica, and Brandie Erhmann at the University of North Carolina at Greensboro for helpful suggestions and comments regarding this work.

Note: Please consult all relevant material safety data sheets (MSDS) before use. Mercury chloride is a toxic chemical. Personal protective equipment (gloves, safety goggles, and lab coat) must be worn when handing it and all associated solutions. Dispose of solutions in clearly labeled chemical waste bottles designated for heavy metals.
1. Preparation of 5 mM Degassed Ammonium Formate Buffer, pH 7.5
<ol>
	<li>Dissolve 0.1576 g of ammonium formate buffer in 450 ml of HPLC grade water. Adjust the pH of the above solution with 1 M formic acid and 1 M ammonium hydroxide to 7.5. Transfer this solution to a 500 ml volumetric flask and add HPLC water to the calibration line to make a 5 mM ammonium formate solution.</li>
	<li>Degas the 5 mM ammonium formate buffer under a vacuum system for 10 min and purge with argon. Repeat twice and store solution under argon. On the day of use, filter the buffer solution through a 0.2 micron filter before use.</li>
</ol>
2. Preparation of Mercury(II) Chloride Solutions
<ol>
	<li>Weigh out 0.2375 g mercury(II) chloride. Dissolve it in 25 ml of 5 mM ammonium formate buffer to produce a 0.035 M mercury(II) chloride solution.</li>
	<li>Add 0.214 ml of 0.035 M mercury(II) chloride solution to 9.785 ml of 5 mM ammonium formate buffer to create a 7.5 x 10-4 M solution. Blanket the 7.5 x 10-4 M mercury(II) solution with argon gas.</li>
</ol>
3. Preparation of CGGC Stock Solution
<ol>
	<li>Dissolve 2.0 mg of the dicysteinyl tetrapeptide, CGGC, in 0.118 ml of HPLC grade acetonitrile and then add 1.0647 ml of 5 mM ammonium formate, pH 7.5 buffer that has been degassed in argon to yield a 5 mM CGGC stock solution.</li>
	<li>Add 225 &#181;l of the 5 mM CGGC stock solution to 1,275 &#181;l of 5 mM ammonium formation pH 7.5 buffer to give a 7.5 x 10-4 M CGGC solution.</li>
</ol>
4. Preparation of Various Reaction Mixtures of Mercury(II) and CGGC
<ol>
	<li>Preparation of 1:0.5 ratio of mercury(II):CGGC solution
	<ol>
		<li>Place 255 &#181;l of 5 mM ammonium formate, pH 7.5 buffer into a 1.5 ml microcentrifuge tube. Add 30 &#181;l of 7.5 x10-4 M mercury(II) chloride solution into the 1.5 ml microcentrifuge tube with the ammonium formate buffer.</li>
		<li>Vortex the solution for 10 sec. Then add 15 &#181;l of 7.5 x 10-4 M CGGC solution into the 1.5 ml microcentrifuge. Vortex the solution for 10 sec. Let the solution stand for 10 min&#160;prior to injection into the mass spectrometer.</li>
	</ol>
	</li>
	<li>Preparation of 1:1 ratio of mercury(II):CGGC solution
	<ol>
		<li>Place 240 &#181;l of 5 mM ammonium formate, pH 7.5 buffer into a 1.5 ml microcentrifuge tube. Add 30 &#181;l of 7.5 x10-4 M mercury(II) chloride solution into the 1.5 ml microcentrifuge tube with the ammonium formate buffer.</li>
		<li>Vortex the solution for 10 sec. Then add 30 &#181;l of 7.5 x 10 -4 M CGGC solution into the 1.5 ml microcentrifuge tube. Repeat in a similar manner as described in section 4.1.</li>
	</ol>
	</li>
	<li>Preparation of 1:2 ratio of mercury(II):CGGC solution
	<ol>
		<li>Place 210 &#181;l of 5 mM ammonium formate, pH 7.5 buffer into a 1.5 ml microcentrifuge tube. Add 30 &#181;l of 7.5 x10-4 M mercury(II) chloride solution into the 1.5 ml microcentrifuge tube with the ammonium formate buffer.</li>
		<li>Vortex the solution for 10 sec. Then add 60 &#181;l of 7.5 x 10-4 M CGGC solution into the 1.5 ml microcentrifuge tube. Repeat in a similar manner as described in section 4.1.</li>
	</ol>
	</li>
</ol>
5. Preparation of CEEC Stock Solution
<ol>
	<li>Dissolve 3.5 mg of the dicysteinyl tetrapeptide, CEEC, in 0.145 ml of HPLC grade acetonitrile to dissolve the peptide. Then add 13.067 ml of 5 mM ammonium formate, pH 7.5 buffer that has been degassed in argon to produce 0.5 mM CEEC solution.</li>
	<li>Vortex the solution until all peptide is dissolved. Add 1.125 ml of 0.5 mM CEEC solution and 0.375 ml of 5 mM ammonium formate, pH 7.5 buffer to a 1.5 ml microcentrifuge tube to give a 7.5 x 10-5 M CEEC solution. Vortex until mixed.</li>
</ol>
6. Preparation of Various Reaction Mixtures of Mercury(II) and CEEC Solution
<ol>
	<li>Preparation of 1:0.5 ratio of mercury(II):CEEC solution
	<ol>
		<li>Place 255 &#181;l of 5 mM ammonium formate, pH 7.5 buffer into a 1.5 ml microcentrifuge tube. Add 30 &#181;l of 7.5 x10-4 M mercury(II) chloride solution into the 1.5 ml microcentrifuge tube with the ammonium formate buffer.</li>
		<li>Vortex the solution for 10 sec. Then add 15 &#181;l of 7.5 x 10-4 M CEEC solution into the 1.5 ml microcentrifuge tube. Repeat in a similar manner as described in section 4.1.</li>
	</ol>
	</li>
	<li>Preparation of 1:1 ratio of mercury(II):CEEC solution
	<ol>
		<li>Place 240 &#181;l of 5 mM ammonium formate, pH 7.5 buffer into a 1.5 ml microcentrifuge tube. Add 30 &#181;l of 7.5 x10-4 M mercury(II) chloride solution into the 1.5 ml microcentrifuge tube with the ammonium formate buffer.</li>
		<li>Vortex the solution for 10 sec. Then add 30 &#181;l of 7.5 x 10-4 M CEEC solution into the 1.5 ml microcentrifuge tube. Repeat in a similar manner as described in section 4.1.</li>
	</ol>
	</li>
	<li>Preparation of 1:2 ratio of mercury(II):CEEC solution
	<ol>
		<li>Place 210 &#181;l of 5 mM ammonium formate, pH 7.5 buffer into a 1.5 ml microcentrifuge tube. Add 30 &#181;l of 7.5 x10-4 M mercury(II) chloride solution into the 1.5 ml microcentrifuge tube with the ammonium formate buffer.</li>
		<li>Vortex the solution for 10 sec. Then add 60 &#181;l of 7.5 x 10-4 M CEEC solution into the 1.5 ml microcentrifuge tube. Repeat in a similar manner as described in section 4.1.</li>
	</ol>
	</li>
</ol>
7. Analyzing the Reaction Mixtures of Mercury(II) and CGGC Samples by Orbitrap ESI Mass Spectrometry
<ol>
	<li>Preparing the ESI mass spectrometer16
	<ol>
		<li>Draw 100 &#181;l of calibration standards into a 500 &#181;l glass syringe.</li>
		<li>Place the syringe in the syringe cradle of the MS pump, attach the tubing, and inject into the mass spectrophotometer.</li>
		<li>Set up the file name for the run by selecting the file icon and typing the file name.</li>
		<li>Select acquire data button in the data acquisition module and collect 150 scans.</li>
		<li>Analyze the chromatogram to verify the calibration standards by opening data processing module of the software. Open the module, go to file menu and select &#34;open&#34;, and select the file in the dialog box. Verify that the peaks in the chromatogram correlate to the mass to charge ratios of the standards.</li>
		<li>Clean the 500 &#181;l glass syringe by drawing up 500 &#181;l HPLC grade methanol and then dispense the methanol into a beaker.</li>
		<li>Draw up 500 &#181;l of HPLC grade methanol into the glass syringe and flush the system as per step 7.1.2.</li>
		<li>Select the method setup module of the software to set the parameters. Choose the scan mode menu and identify the analyzer as FTMS, and then click on &#34;OK&#34;. Then by clicking on the various icons on the real-time view spectrum page, set the following parameters: Sheath gas flow rate: 10, Source temperature: 0, Capillary voltage: 37 V, Tube lens: 95 V, Spray voltage: 4.20 kV, Flow rate 10.00 &#181;l/min, Analyzer: FTMS, Number of scans: 150.</li>
	</ol>
	</li>
	<li>Running CGGC samples on ESI mass spectrometer
	<ol>
		<li>Run the 5 mM ammonium formate pH 7.5 buffer.
		<ol>
			<li>Place 500 &#181;l of 5 mM ammonium formate buffer into the 500 &#181;l glass syringe, place it into the syringe cradle of the MS pump, and attach the tubing.</li>
			<li>Run buffer through the tubing for 1-2 min.</li>
			<li>Set up the file name for the run by selecting the file icon and typing the file name.</li>
			<li>Select acquire data button in the module and collect 150 scans.</li>
			<li>Click the run button to stop collection after 150 scans are collected.</li>
			<li>Open the data browser module, then go to file menu and select &#34;open&#34;, and select the file in the dialog box. Verify that no peaks at 483, 683, 1,163 and 1,363 m/z are present that resemble the peptide or mercury (II)-peptide complexes.</li>
		</ol>
		</li>
		<li>Run the 1:0.5 mercury(II):CGGC ratio solution.
		<ol>
			<li>Place 250 &#181;l of 1:0.5 mercury(II):CGGC ratio of the sample into the syringe.</li>
			<li>Place the syringe into the syringe cradle of the MS pump, attach the tubing, and prime the apparatus.</li>
			<li>Select a file name for the run by selecting the file icon and typing the file name.</li>
			<li>Press the acquire data button in the data acquisition module and collect 150 scans and click the run button to stop the collection.</li>
			<li>Open the module, go to file menu and select &#34;open&#34;, and select the file in the dialog box. Verify that the chromatogram contains peaks including the one for the CGGC peptide alone.</li>
			<li>Wash the syringe by aspirating with 500 &#181;l ammonium formate buffer and then dispensing the ammonium formate buffer into a beaker.</li>
			<li>Select the waste button on the MS and flush the tubing three times with 500 &#181;l ammonium formate buffer.</li>
			<li>Wash the syringe by aspirating with 500 &#181;l methanol and then dispensing the methanol into a beaker.</li>
			<li>Flush the tubing one time with 500 &#181;l methanol.</li>
			<li>Select load detector button on the MS.</li>
			<li>Add 500 &#181;l of ammonium formate buffer to the syringe.</li>
			<li>Place the syringe into the syringe cradle of the MS pump, attach the tubing, and prime the apparatus.</li>
			<li>Select a file name for the buffer run by selecting the file icon and typing the file name.</li>
			<li>Press the acquire data button and collect 150 scans and then click the stop run button.</li>
			<li>Open the data browser module, go to file menu and select &#34;open&#34;, and select the file in the dialog box. Verify that the chromatogram is void of peaks from the previous Hg:CGGC run.</li>
		</ol>
		</li>
		<li>Run the 1:1 mercury(II):CGGC ratio solution.
		<ol>
			<li>Place 250 &#181;l of 1:1 mercury(II):CGGC ratio sample into the syringe.</li>
			<li>Repeat in a similar manner as described for Steps 7.2.2.2 to 7.2.2.15.</li>
		</ol>
		</li>
		<li>Run the 1:2 mercury(II):CGGC ratio solution
		<ol>
			<li>Place 250 &#181;l of 1:2 mercury(II):CGGC concentration sample into the syringe.</li>
			<li>Repeat in a similar manner as described for Steps 7.2.2.2 to 7.2.2.15.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
8. Analyzing the Reaction Mixtures of Mercury and CEEC Samples by Orbitrap ESI Mass Spectrometry
<ol>
	<li>Running CEEC samples on ESI mass spectrometer
	<ol>
		<li>Repeat analysis procedure (Steps 7.1 to 7.2) using CEEC samples and reaction mixtures of mercury(II) and CEEC at various stoichiometric ratios.</li>
	</ol>
	</li>
</ol>

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<li>Ngu-Schwemlein, M., Lin, X., Rudd, B., Bronson, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Synthesis+and+ESI+mass+spectrometric+analysis+of+the+association+of+mercury%28II%29+with+multi-cysteinyl+peptides%5BTitle%5D)+AND+%22J.+Inorg.+Biochem%22%5BJournal%5D)">Synthesis and ESI mass spectrometric analysis of the association of mercury(II) with multi-cysteinyl peptides.</a> J. Inorg. Biochem. 133, 8-23 (2014).</li>
<li>Winther, J. R., Thorpe, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Quantification+of+thiols+and+disulfides%5BTitle%5D)+AND+%22Biochimica+et.+Biophysica+Acta%22%5BJournal%5D)">Quantification of thiols and disulfides.</a> Biochimica et. Biophysica Acta. 1840, 838-846 (2014).</li>
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<li>Schaumlöffel, D., Tholey, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Recent+directions+of+electrospray+mass+spectrometry+for+elemental+speciation+analysis%5BTitle%5D)+AND+%22Anal.+Bioanal.+Chem%22%5BJournal%5D)">Recent directions of electrospray mass spectrometry for elemental speciation analysis.</a> Anal. Bioanal. Chem. 400, 1645-1652 (2011).</li>
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There is no disclosure for the work reported.

The hydrophobic dicysteinyl tetrapeptide CGGC (C10H18N4O5S2; MW = 338) (Figure 1), forms complexes with mercury(II) as shown in Figure 2 and Table 1. Additionally, it forms peptide dimers and trimers incrementally as the amount of peptide increases in the reaction mixture. As shown by the m/z values of the associated dimers [(2M+H)+ = 677] and trimers [(3M+H)+ = 1015], the thiol groups of CGGC did not oxidize to form disulfides under the experimental conditions. The formation of these associated CGGC species could be due to the hydrophobicity of this tetrapeptide. CGGC forms two types of complexes with mercury corresponding to 1:1 mercury(II):peptide and 1:2 mercury (II):(peptide)2 complexes as previously reported for dicysteinyl tripeptides7. However, in the presence of excess or equivalent mercury(II), it also forms a 2:2 [mercury(II)]2:(peptide)2 complex.
The carboxylated dicysteinyl tetrapeptide CEEC (C16H26N4O9S2; MW = 482) (Figure 1) form complexes with mercury(II) as shown in Figure 3 and Table 1. It did not form CEEC dimers as readily as that observe for the more hydrophobic CGGC. Comparable to CGGC, it forms complexes with mercury corresponding to 1:1 mercury(II):peptide and 1:2 mercury (II):(peptide)2 complexes. However, with the auxiliary carboxylate groups, it forms the 2:2 [mercury(II)]2:(peptide)2 complex more readily. Moreover, in excess mercury, it forms the 2:1 [mercury(II)]2:peptide complex and the 4:2 [mercury(II)]4:(peptide)2 peptide complex, which were not observed for CGGC.
The summary of the observed signals for the complexes formed as m/z values are shown in Table 1.
<img alt="Table 1" src="/files/ftp_upload/53536/53536table1.jpg" />
Table 1. Summary of mercury-peptide complexes signals. Mercury-peptide complexes signals in the LTQ/Orbitrap MS chromatograms in ammonium formate buffer, pH 7.5.
We have demonstrated that the reaction of mercury(II) and two dicysteinyl tetrapeptides form complexes that are dependent on the initial ratios of mercury(II):peptide as well as the presence of auxiliary binding groups in the dicysteinyl tetrapeptide. Moreover, accurate stoichiometry of mercury and peptide in the complexes formed under specified electrospray ionization conditions can be determined by using high resolution ESI mass spectrometry based on distinct mercury isotopic distribution patterns.
In reacting cysteinyl peptides with mercury(II), precautions must be taken to prevent the oxidation of cysteinyl thiol groups to form disulfide bonds. Within the described protocol, the buffer solutions were carefully degassed and stored under argon. In addition, all reaction samples are prepared immediately before analysis by ESI mass spectrometry.
Due to differences in solubility between the two tetrapeptides, CEEC and CGGC, different concentrations were used to prepare the stock solutions. The freezer stock of CGGC peptide was wetted with acetonitrile and was easily dissolved followed by 5 mM ammonium formate buffer, pH 7.5 to produce a 7.5 x 10-4 M CGGC solution. The CEEC was prepared at a lower concentration, 7.5 x 10-5 M, prior to the mercury(II):peptide reaction mixture steps because of its lower solubility. The optimal dilution for analyzing the mercury(II) complexes was deemed to be 10-5 M because of the solubility of the peptide and to allow for removal of residues in the mass spectrometer. In contract to the CGGC solutions, CEEC residues adhere to the tubing, which necessitates occasional tubing replacement.
The significance of using ESI mass spectrometry for the analysis of mercury-peptide complexes lies in its soft ionization of analytes. This facilitates the analysis of molecular ions with negligible fragmentation. As shown in this work, it can be used to characterize the stoichiometries of mercury-peptide complexes based on the signature mercury isotopic distribution patterns. However, a volatile buffer system is necessary for analysis by ESI mass spectrometry. This may limit its practical use for identifying analytes that require less volatile solvents or buffering media for dissolution.
As we have previously mentioned7,8, ESI mass spectrometry provides a sensitive analytical tool for an accurate determination of the stoichiometry of mercury and peptide in the mercury-peptide complexes under the specified electrospray ionization condition. However, it is necessary to use additional methods (for example, 1H, 13C, 199Hg NMR spectroscopy, extended X-ray absorption fine structure, or potentiometry17-18) to provide a more accurate determination of the content of complexes in solution.
We have shown that ESI with an orbitrap mass analyzer can be used to analyze mercury-peptide complexes. We expect that this technique can be applied toward the analysis of other metal ions and their complexes with various small compounds. It will be especially useful for analyzing complexes formed by other metal ions that can exist in different isotopic forms.

Current clinical drugs prescribed for chelation therapy of mercury poisoning1 contain thiol group(s), which is/are responsible for binding and sequestering mercury ions2,3. However, studies have shown that these small thiol compounds [dimercaptosuccinic acid (DMSA) and dimercaptopropane-sulfonic acid (DMPS)] are not optimal for mercury chelation therapy4-6. Therefore, there is a need to understand the association and complex formation tendencies of mercury with thiols to enhance the rational drug design of thiol compounds for mercury chelation. Recently, we reported that n-alkyl and aryl dicysteinyl tripeptides with dithiol groups can serve as effective "double anchors" to accommodate the coordination sites of mercury(II) to form 1:1 mercury(II):peptide and 1:2 mercury(II):(peptide)2 complexes7. Additionally, we studied the effect of increasing cysteinyl residues on complex type formations8. In this study, we investigate the association of mercury(II) with two dicysteinyl tetrapeptides, where the cysteinyl residues are separated by two amino acid residues. In order to evaluate the effect of auxiliary binding groups for mercury, the intervening amino acids are either two glycine (unsubstituted) residues or two glutamic acid (gamma-carboxylated) residues. 
The reaction of cysteinyl peptide with mercury(II) requires conditions that will prevent the oxidation of the cysteinyl thiol groups to form disulfide bonds9. Moreover, the association of mercury(II) with cysteinyl peptides to form various types of mercury-peptide complexes is dependent on the initial ratio of mercury(II): peptide in the reaction mixture7,8. The types of mercury-peptide complexes formed in these reaction mixtures can be analyzed by soft-ionization mass spectroscopy, which is a sensitive analytical tool for determining species interactions between metal ions and peptides10-14. Accordingly, it will provide a profile of the various types of mercuriated peptide adducts that are formed under a specified electrospray ionization condition. Here, we will show how cysteinyl peptides and mercury(II) chloride solutions can be prepared in degassed ammonium formate buffer solution blanketed with argon to minimize oxidation. By reacting varying mole equivalents of mercury(II) with dicysteinyl tetrapeptides, we will show how the initial ratio of mercury(II):peptide has an effect on the types of complexes formed. We will also show how electrospray ionization (ESI) mass spectrometry can be used as a characterization tool to provide an accurate stoichiometry of mercury to peptide in the complexes formed. The associated video protocol will demonstrate the experimental conditions for preparing the mercury complexes, the procedure for analyzing the reaction mixtures under specified electrospray ionization conditions, and the characterization of the stoichiometries of mercury(II)-dicysteinyl tetrapeptide complexes, based on the distinct mercury isotope distribution patterns, by using the ChemCal program15. It is intended to assist those who are interested in using ESI orbitrap mass spectrometry to analyze various complexes formed by metal ions that exist in different isotopic forms.

<table><tbody><tr><td>Mercury(II) chloride</td><td>Sigma-Aldrich</td><td>429724</td><td>Highly toxic</td></tr><tr><td>Ammonium formate</td><td>Sigma-Aldrich</td><td>516961</td><td></td></tr><tr><td>Formic acid</td><td>Sigma-Aldrich</td><td>F0507</td><td></td></tr><tr><td>Ammonium hydroxide</td><td>Fisher</td><td>A512-P500</td><td></td></tr><tr><td>HPLC water</td><td>Fisher</td><td>W5-4</td><td></td></tr><tr><td>HPLC Acetonitrile</td><td>Fisher</td><td>BP2405-1</td><td></td></tr><tr><td>HPLC Methanol</td><td>Fisher</td><td>A452-4</td><td></td></tr></tbody></table>

a study of the complexation of mercury(ii) with dicysteinyl tetrapeptides by electrospray ionization mass spectrometry

In this study we evaluated a method for the characterization of complexes, formed in different relative ratios of mercury(II) to dicysteinyl tetrapeptide, by electrospray ionization orbitrap mass spectrometry. This strategy is based on previous successful characterization of mercury-dicysteinyl complexes involving tripeptides by utilizing mass spectrometry among other techniques. Mercury(II) chloride and a dicysteinyl tetrapeptide were incubated in a degassed buffered medium at varying stoichiometric ratios. The complexes formed were subsequently analyzed on an electrospray mass spectrometer consisting of a hybrid linear ion- and orbi- trap mass analyzer. The electrospray ionization mass spectrometry (ESI-MS) spectra were acquired in the positive mode and the observed peaks were then analyzed for distinct mercury isotopic distribution patterns and associated monoisotopic peak. This work demonstrates that an accurate stoichiometry of mercury and peptide in the complexes formed under specified electrospray ionization conditions can be determined by using high resolution ESI MS based on distinct mercury isotopic distribution patterns.

A study was performed to characterize the possible mercury-peptide complex composition for two tetrapeptides, CGGC and CEEC (Figure 1) by ESI mass spectrometry. Complexes of mercury(II) with CGGC or CEEC were investigated by reacting the mixtures of mercury(II) and peptide solutions at three different molar ratios: 1:0.5, 1:1, and 1:2 (mercury(II): peptide). The concentration of mercury(II) was 7.5 x 10-6 M and the peptide concentration varied accordingly.
<img alt="Figure 1" src="/files/ftp_upload/53536/53536fig1.jpg" /> 
Figure 1. Dicysteinyl peptide structures. Chemical structures of the dicysteinyl tetrapeptides, CGGC and CEEC. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53536/53536fig2.jpg" /> 
Figure 2. ESI MS of mercury(II) and CGGC. Electrospray ionization orbitrap mass spectra from a solution containing 7.5 x 10-6 M Hg2+ in ammonium formate buffer, pH 7.5 containing varying Hg2+: CGGC stoichiometric ratios: (A) 1:0.5 ratio, (B) 1:1 ratio, and (C) 1:2 ratio. Insets show the mercury isotopic patterns of the indicated mercury-peptide complexes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53536/53536fig3.jpg" /> 
Figure 3. ESI MS of mercury(II) and CEEC. Electrospray ionization orbitrap mass spectra from a solution containing 7.5 x 10-6 M Hg2+ in ammonium formate buffer, pH 7.5 containing varying Hg2+: CEEC stoichiometric ratios: (A) 1:0.5 ratio, (B) 1:1 ratio, and (C) 1:2 ratio. Insets show the mercury isotopic patterns of the indicated mercury-peptide complexes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Electrospray ionization orbitrap mass chromatograms were collected for mercury(II) complexing with CGGC (Figure 2) and CEEC (Figure 3) at various mercury(II) to peptide stoichiometric ratios (1:0.5, 1:1, and 1:2). The observed mercury-peptide complex types show distinct mercury isotopic peaks (insets), which are used to determine the number of mercury ions in the complex as well as the number of deprotonations. For example, Figure 1b inset shows the mercury isotopic signature in the peptide-mercury adduct, which corresponds to the seven main naturally occurring isotopes of mercury: 196Hg (0.146%), 198Hg (10.02%), 199Hg (16.84%), 200Hg (23.13%), 201Hg (13.22%), 202Hg (29.80%), 204Hg (6.85%), with percent natural abundances indicated in parentheses. The two major isotopes 200Hg and 202Hg show a distinct relative intensity ratio of 2.3:3. Accordingly the most intense isotopic peak of this one-mercury isotope cluster constitutes the monoisotopic mass for the adduct (m/z = 539). It correlates with a two-coordinate complex, which is formed by the deprotonation of two cysteinyl thiols to form the [(CGGC-2H+Hg)+H]+ adduct. This analysis is made as follows:
m/z value for [(CGGC-2H+Hg)+H]+ is equal to (338 - 2 + 202 +1) = 539.
Figure 1A inset shows the mercury isotopic signature in the peptide-mercury adduct, which corresponds to a two-mercury complex as calculated by using the ChemCal program for [(2CGGC-4H+2Hg)+H]+ (Figure 4). The theoretical protonated monoisotopic mass corresponds to an m/z value of 1077.061, which is the ninth isotopic peak in the calculated isotopic cluster. Figure 1A inset shows an isotopic peak corresponding to an m/z value of 1077.1, which is also the ninth peak in the observed isotopic cluster. Therefore, the originating adduct for this isotopic cluster can be assigned for [(2CGGC-4H+2Hg)+H]+.
<img alt="Figure 4" src="/files/ftp_upload/53536/53536fig4.jpg" /> 
Figure 4. Theoretical isotopic patterns for [(2CGGC-4H+2Hg)+H]+. The theoretical isotopic patterns for [(2CGGC-4H+2Hg)+H]+ as calculated by using the ChemCal program. Arrow indicates monoisotopic peak. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/53536/53536fig5.jpg" /> 
Figure 5. Cationized adducts. Some cationized sodium and potassium adducts associated with the mercury-peptide complexes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Figure 5 shows some cationized sodium and potassium adducts associated with the mercury-peptide complexes formed by CGGC. Sodiated adducts are 22 mass units larger than the corresponding protonated mercury-CGGC complexes, whereas the potassium adducts are 38 mass units larger. The dominant protonated CGGC dimer (m/z = 677) also forms cationized species with sodium (m/z = 699) and potassium ions (m/z = 715). This further confirms the formation of CGGC dimers without the oxidation of the cysteinyl thiol groups to form disulfides, which would have resulted in a decrease of two mass units for protonated or cationized adducts.
<img alt="Figure 6" src="/files/ftp_upload/53536/53536fig6.jpg" /> 
Figure 6. Overlapping +1 and +2 charge states. Overlapping peaks associated with mercury-peptide ions [(CEEC-4H+2Hg)+H]+ in the +1 and +2 charge states. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/53536/53536fig7.jpg" /> 
Figure 7. Theoretical isotopic patterns for [(CEEC-4H+2Hg)+H]+. The theoretical isotopic patterns for [(CEEC-4H+2Hg)+H]+ as calculated by using the ChemCal program. Arrow indicates monoisotopic peak. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Figure 6 shows overlapping peaks associated with mercury-CEEC adducts in the +1 and +2 charge. It shows isotopic peaks that are associated with mercury-peptide ions [(CEEC-4H+2Hg)+H]+ in the +1 charge and an m/z value of 883. This is in agreement with a two mercury complex as calculated for [(CEEC-4H+2Hg)+H]+ by using the ChemCal program (Figure 7). The theoretical protonated monoisotopic mass corresponds to an m/z value of 883.032.
The above observed [(CEEC-4H+2Hg)+H]+ adduct with a monoisotopic peak of 883.03 overlaps with another adduct containing corresponding peaks showing an additional 0.5 mass units. With the extremely high resolution achieved by the orbitrap mass spectrometry instrument, it can be postulated that these overlapping peaks correspond to adducts with a charge of +2. Accordingly, the monoisotopic mass of the overlapping complex being ionized can be calculated as follows. Figure 8 shows that the m/z difference between the isotopic peaks is 0.5 and the mass difference between them is 1 amu. Therefore, the charge state is +2. To calculate the mass of the mercury-peptide complex, the m/z for the monoisotopic peak is multiplied by the charge state, and subtracted from the mass of two protons, which made the complex ion positively charged.
Calculations for the +2 adduct:
m/z difference between isotopic peaks is 0.5
Mass difference between isotopic peaks is 1 amu (1 neutron)
z = 1 divide by 0.5 = 2
m/z for protonated monoisotopic peak is (883.53 x 2) - 2 = 1765.06
The above m/z value for the protonated monoisotopic peak, [(2CEEC-8H+4Hg)+H]+, is consistent with the theoretical value as calculated by the ChemCal program as 1765.056 (Figure 8).
<img alt="Figure 8" src="/files/ftp_upload/53536/53536fig8.jpg" /> 
Figure 8. Theoretical isotopic patterns for [(2CEEC-8H+4Hg)+H]+. The theoretical isotopic patterns for [(2CEEC-8H+4Hg)+H]+ as calculated by using the ChemCal program. Arrow indicates monoisotopic peak. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53536/53536fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The advantage of analyzing mercury-peptide complexes with an ESI orbitrap mass spectrometer is that the charge of every ion can be readily assigned as shown above. Peptides containing basic amino-terminus can readily stabilize positive charges. When using electrospray ionization and a high resolution mass analyzer such as the orbitrap, the charge state of peptide ions with greater than +1 charge can be determined more readily compared to the lower resolution iontrap mass analyzer.
The overlapping peaks associated with mercury-CEEC adducts (Figure 3A and Figure 6), as described above, were also analyzed by tandem MS. It did not show any MS-MS fragmentation, which indicated that the obtained signals belong to the expected compound as discussed above, and are not clustered artifacts formed at higher concentrations of mercury-to-peptide ratios.

Watch this Scientific Journal Video about A Study of the Complexation of MercuryII with Dicysteinyl Tetrapeptides by Electrospray Ionization Mass Spectrometry at JoVE.com

A Study of the Complexation of MercuryII with Dicysteinyl Tetrapeptides by Electrospray Ionization Mass Spectrometry

The characterization of complexes formed in different relative ratios of mercury(II) to dicysteinyl tetrapeptides by electrospray ionization orbitrap mass spectrometry is presented.

A Study of the Complexation of Mercury(II) with Dicysteinyl Tetrapeptides by Electrospray Ionization Mass Spectrometry

In this study we evaluated a method for the characterization of complexes, formed in different relative ratios of mercury(II) to dicysteinyl tetrapeptide, ...

a-study-complexation-mercury-ii-with-dicysteinyl-tetrapeptides

Nanyang Technological University

Research

JoVE Journal

Chemistry

7.9K Views.  University of North Carolina at Greensboro. The characterization of complexes formed in different relative ratios of mercury(II) to dicysteinyl tetrapeptides by electrospray ionization orbitrap mass spectrometry is presented.

Video: A Study of the Complexation of MercuryII with Dicysteinyl Tetrapeptides by Electrospray Ionization Mass Spectrometry

Production of Disulfide-stabilized Transmembrane Peptide Complexes for Structural Studies

Physical interactions among the lipid-embedded alpha-helical domains of membrane proteins play a crucial role in folding and assembly of membrane protein complexes and in dynamic processes such as transmembrane (TM) signaling and regulation of cell-surface protein levels. Understanding the structural features driving the association of particular sequences requires sophisticated biophysical and biochemical analyses of TM peptide complexes. However, the extreme hydrophobicity of TM domains makes them very difficult to manipulate using standard peptide chemistry techniques, and production of suitable study material often proves prohibitively challenging. Identifying conditions under which peptides can adopt stable helical conformations and form complexes spontaneously adds a further level of difficulty. Here we present a procedure for the production of homo- or hetero-dimeric TM peptide complexes from materials that are expressed in E. coli, thus allowing incorporation of stable isotope labels for nuclear magnetic resonance (NMR) or non-natural amino acids for other applications relatively inexpensively. The key innovation in this method is that TM complexes are produced and purified as covalently associated (disulfide-crosslinked) assemblies that can form stable, stoichiometric and homogeneous structures when reconstituted into detergent, lipid or other membrane-mimetic materials. We also present carefully optimized procedures for expression and purification that are equally applicable whether producing single TM domains or crosslinked complexes and provide advice for adapting these methods to new TM sequences.

Biophysical and biochemical studies of interactions among membrane-embedded protein domains face many technical challenges, the first of which is obtaining appropriate study material. This article describes a protocol for producing and purifying disulfide-stabilized transmembrane peptide complexes that are suitable for structural analysis by solution nuclear magnetic resonance (NMR) and other analytical applications.

Physical  interactions among the lipid-embedded alpha-helical domains of membrane  proteins play a crucial role in folding and assembly of membrane ...

Compact Quantum Dots for Single-molecule Imaging

Single-molecule imaging is an important tool for understanding the mechanisms of biomolecular function and for visualizing the spatial and temporal heterogeneity of molecular behaviors that underlie cellular biology 1-4. To image an individual molecule of interest, it is typically conjugated to a fluorescent tag (dye, protein, bead, or quantum dot) and observed with epifluorescence or total internal reflection fluorescence (TIRF) microscopy. While dyes and fluorescent proteins have been the mainstay of fluorescence imaging for decades, their fluorescence is unstable under high photon fluxes necessary to observe individual molecules, yielding only a few seconds of observation before complete loss of signal. Latex beads and dye-labeled beads provide improved signal stability but at the expense of drastically larger hydrodynamic size, which can deleteriously alter the diffusion and behavior of the molecule under study.
Quantum dots (QDs) offer a balance between these two problematic regimes. These nanoparticles are composed of semiconductor materials and can be engineered with a hydrodynamically compact size with exceptional resistance to photodegradation 5. Thus in recent years QDs have been instrumental in enabling long-term observation of complex macromolecular behavior on the single molecule level. However these particles have still been found to exhibit impaired diffusion in crowded molecular environments such as the cellular cytoplasm and the neuronal synaptic cleft, where their sizes are still too large 4,6,7.
Recently we have engineered the cores and surface coatings of QDs for minimized hydrodynamic size, while balancing offsets to colloidal stability, photostability, brightness, and nonspecific binding that have hindered the utility of compact QDs in the past 8,9. The goal of this article is to demonstrate the synthesis, modification, and characterization of these optimized nanocrystals, composed of an alloyed HgxCd1-xSe core coated with an insulating CdyZn1-yS shell, further coated with a multidentate polymer ligand modified with short polyethylene glycol (PEG) chains (Figure 1). Compared with conventional CdSe nanocrystals, HgxCd1-xSe alloys offer greater quantum yields of fluorescence, fluorescence at red and near-infrared wavelengths for enhanced signal-to-noise in cells, and excitation at non-cytotoxic visible wavelengths. Multidentate polymer coatings bind to the nanocrystal surface in a closed and flat conformation to minimize hydrodynamic size, and PEG neutralizes the surface charge to minimize nonspecific binding to cells and biomolecules. The end result is a brightly fluorescent nanocrystal with emission between 550-800 nm and a total hydrodynamic size near 12 nm. This is in the same size range as many soluble globular proteins in cells, and substantially smaller than conventional PEGylated QDs (25-35 nm).

We describe the preparation of colloidal quantum dots with minimized hydrodynamic size for single-molecule fluorescence imaging. Compared to conventional quantum dots, these nanoparticles are similar in size to globular proteins and are optimized for single-molecule brightness, stability against photodegradation, and resistance to nonspecific binding to proteins and cells.

Single-molecule imaging is an important tool for  understanding the mechanisms of biomolecular function and for visualizing the  spatial and temporal ...

Engineering

Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique amino acid sequence, it is only realized by the folding of the polypeptide chain into a single defined three-dimensional arrangement or more commonly into an ensemble of interconverting conformations. Investigating the connection between protein conformation and its function is therefore essential for a complete understanding of how proteins are able to fulfill their great variety of tasks. One possibility to study conformational changes a protein undergoes while progressing through its functional cycle is hydrogen-1H/2H-exchange in combination with high-resolution mass spectrometry (HX-MS). HX-MS is a versatile and robust method that adds a new dimension to structural information obtained by e.g. crystallography. It is used to study protein folding and unfolding, binding of small molecule ligands, protein-protein interactions, conformational changes linked to enzyme catalysis, and allostery. In addition, HX-MS is often used when the amount of protein is very limited or crystallization of the protein is not feasible. Here we provide a general protocol for studying protein dynamics with HX-MS and describe as an example how to reveal the interaction interface of two proteins in a complex. &#160;&#160;

Protein conformation and dynamics are key to understanding the relationship between protein structure and function. Hydrogen exchange coupled with high-resolution mass spectrometry is a versatile method to study the conformational dynamics of proteins as well as characterizing protein-ligand and protein-protein interactions, including contact interfaces and allosteric effects.

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique ...

Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. Hydrogen-Deuterium Exchange (HDX) measured at rapid time scales is uniquely suited to detect structures and hydrogen bonding networks that are briefly populated, allowing for the characterization of transient conformers in native ensembles. Coupling of HDX to mass spectrometry offers several key advantages, including high sensitivity, low sample consumption and no restriction on protein size. This technique has advanced greatly in the last several decades, including the ability to monitor HDX labeling times on the millisecond time scale. In addition, by incorporating the HDX workflow onto a microfluidic platform housing an acidic protease microreactor, we are able to localize dynamic properties at the peptide level. In this study, Time-Resolved ElectroSpray Ionization Mass Spectrometry (TRESI-MS) coupled to HDX was used to provide a detailed picture of residual structure in the tau protein, as well as the conformational shifts induced upon hyperphosphorylation.

Conformational flexibility plays a critical role in protein function. Herein, we describe the use of time-resolved electrospray ionization mass spectrometry coupled to hydrogen-deuterium exchange for probing the rapid structural changes that drive function in ordered and disordered proteins.

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. ...

Biochemistry

Mass Spectrometric Approaches to Study Protein Structure and Interactions in Lyophilized Powders

Amide hydrogen/deuterium exchange (ssHDX-MS) and side-chain photolytic labeling (ssPL-MS) followed by mass spectrometric analysis can be valuable for characterizing lyophilized formulations of protein therapeutics. Labeling followed by suitable proteolytic digestion allows the protein structure and interactions to be mapped with peptide-level resolution. Since the protein structural elements are stabilized by a network of chemical bonds from the main-chains and side-chains of amino acids, specific labeling of atoms in the amino acid residues provides insight into the structure and conformation of the protein. In contrast to routine methods used to study proteins in lyophilized solids (e.g., FTIR), ssHDX-MS and ssPL-MS provide quantitative and site-specific information. The extent of deuterium incorporation and kinetic parameters can be related to rapidly and slowly exchanging amide pools (Nfast, Nslow) and directly reflects the degree of protein folding and structure in lyophilized formulations. Stable photolytic labeling does not undergo back-exchange, an advantage over ssHDX-MS. Here, we provide detailed protocols for both ssHDX-MS and ssPL-MS, using myoglobin (Mb) as a model protein in lyophilized formulations containing either trehalose or sorbitol.

Here, we present detailed protocols for solid-state amide hydrogen/deuterium exchange mass spectrometry (ssHDX-MS) and solid-state photolytic labeling mass spectrometry (ssPL-MS) for proteins in solid powders. The methods provide high-resolution information on protein conformation and interactions in the amorphous solid-state, which may be useful in formulation design.

Amide hydrogen/deuterium exchange (ssHDX-MS) and side-chain photolytic labeling (ssPL-MS) followed by mass spectrometric analysis can be valuable for ...

Ion Mobility-Mass Spectrometry Techniques for Determining the Structure and Mechanisms of Metal Ion Recognition and Redox Activity of Metal Binding Oligopeptides

Electrospray ionization (ESI) can transfer an aqueous-phase peptide or peptide complex to the gas-phase while conserving its mass, overall charge, metal-binding interactions, and conformational shape. Coupling ESI with ion mobility-mass spectrometry (IM-MS) provides an instrumental technique that allows for simultaneous measurement of a peptide&#8217;s mass-to-charge (m/z) and collision cross section (CCS) that relate to its stoichiometry, protonation state, and conformational shape. The overall charge of a peptide complex is controlled by the protonation of 1) the peptide&#8217;s acidic and basic sites and 2) the oxidation state of the metal ion(s). Therefore, the overall charge state of a complex is a function of the pH of the solution that affects the peptides metal ion binding affinity. For ESI-IM-MS analyses, peptide and metal ions solutions are prepared from aqueous-only solutions, with the pH adjusted with dilute aqueous acetic acid or ammonium hydroxide. This allows for pH dependence and metal ion selectivity to be determined for a specific peptide. Furthermore, the m/z and CCS of a peptide complex can be used with B3LYP/LanL2DZ molecular modeling to discern binding sites of the metal ion coordination and tertiary structure of the complex. The results show how ESI-IM-MS can characterize the selective chelating performance of a set of alternative methanobactin peptides and compare them to the copper-binding peptide methanobactin.

Ion mobility-mass spectrometry and molecular modeling techniques can characterize the selective metal chelating performance of designed metal-binding peptides and the copper-binding peptide methanobactin. Developing new classes of metal chelating peptides will help lead to therapeutics for diseases associated with metal ion misbalance.

Electrospray ionization (ESI) can transfer an aqueous-phase peptide or peptide complex to the gas-phase while conserving its mass, overall charge, ...

Analysis of Complex Molecules and Their Reactions on Surfaces by Means of Cluster-Induced Desorption/Ionization Mass Spectrometry

Desorption/Ionization Induced by Neutral SO2 Clusters (DINeC) is employed as a very soft and efficient desorption/ionization technique for mass spectrometry (MS) of complex molecules and their reactions on surfaces. DINeC is based on a beam of SO2 clusters impacting on the sample surface at low cluster energy. During cluster-surface impact, some of the surface molecules are desorbed and ionized via dissolvation in the impacting cluster; as a result of this dissolvation-mediated desorption mechanism, low cluster energy is sufficient and the desorption process is extremely soft. Both surface adsorbates and molecules of which the surface is composed of can be analyzed. Clear and fragmentation-free spectra from complex molecules such as peptides and proteins are obtained. DINeC does not require any special sample preparation, in particular no matrix has to be applied. The method yields quantitative information on the composition of the samples; molecules at a surface coverage as low as 0.1 % of a monolayer can be detected. Surface reactions such as H/D exchange or thermal decomposition can be observed in real-time and the kinetics of the reactions can be deduced. Using a pulsed nozzle for cluster beam generation, DINeC can be efficiently combined with ion trap mass spectrometry. The matrix-free and soft nature of the DINeC process in combination with the MSn capabilities of the ion trap allows for very detailed and unambiguous analysis of the chemical composition of complex organic samples and organic adsorbates on surfaces.

Neutral SO2 clusters of low kinetic energy (&#60; 0.8 eV/constituent) are used to desorb complex surface molecules such as peptides or lipids for further analysis by means of mass spectrometry using an ion trap mass spectrometer. No special sample preparation is required, and real-time observation of reactions is possible.

Desorption/Ionization Induced by Neutral SO2 Clusters (DINeC) is employed as a very soft and efficient desorption/ionization technique for mass ...

Capillary Electrophoresis-based Hydrogen/Deuterium Exchange for Conformational Characterization of Proteins with Top-down Mass Spectrometry

Resolving conformational heterogeneity of multiple protein states that coexist in solution remains one of the main obstacles in the characterization of protein therapeutics and the determination of the conformational transition pathways critical for biological functions, ranging from molecular recognition to enzymatic catalysis. Hydrogen/deuterium exchange (HDX) reaction coupled with top-down mass spectrometric (MS) analysis provides a means to characterize protein higher-order structures and dynamics in a conformer-specific manner. The conformational resolving power of this technique is highly dependent on the efficiencies of separating protein states at the intact protein level and minimizing the residual non-deuterated protic content during the HDX reactions. 
Here we describe a capillary electrophoresis (CE)-based variant of the HDX MS approach that aims to improve the conformational resolution. In this approach, proteins undergo HDX reactions while migrating through a deuterated background electrolyte solution (BGE) during the capillary electrophoretic separation. Different protein states or proteoforms that coexist in solution can be efficiently separated based on their differing charge-to-size ratios. The difference in electrophoretic mobility between proteins and protic solvent molecules minimizes the residual non-deuterated solvent, resulting in a nearly complete deuterating environment during the HDX process. The flow-through microvial CE-MS interface allows efficient electrospray ionization of the eluted protein species following a rapid mixing with the quenching and denaturing modifier solution at the outlet of the sprayer. The online top-down MS analysis measures the global deuteration level of the eluted intact protein species, and subsequently, the deuteration of their gas-phase fragments. This paper demonstrates this approach in differential HDX for systems, including the natural protein variants coexisting in milk.

Presented here is a protocol for a capillary electrophoresis-based hydrogen/deuterium exchange (HDX) approach coupled with top-down mass spectrometry. This approach characterizes the difference in higher-order structures between different protein species, including proteins in different states and different proteoforms, by conducting concurrent differential HDX and electrophoretic separation.

Resolving conformational heterogeneity of multiple protein states that coexist in solution remains one of the main obstacles in the characterization of ...

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 ...

Biology

Thermochemical Studies of Ni(II) and Zn(II) Ternary Complexes Using Ion Mobility-Mass Spectrometry

This article describes an experimental protocol using electrospray-ion mobility-mass spectrometry (ES-IM-MS) and energy-resolved threshold collision-induced dissociation (TCID) to measure the thermochemistry of the dissociation of negatively-charged [amb+M(II)+NTA]- ternary complexes into two product channels: [amb+M(II)] + NTA or [NTA+M(II)]-&#160;+ amb, where M = Zn or Ni and NTA is nitrilotriacetic acid. The complexes contain one of the alternative metal binding (amb) heptapeptides with the primary structures acetyl-His1-Cys2-Gly3-Pro4-Tyr5-His6-Cys7 or acetyl-Asp1-Cys2-Gly3-Pro4-Tyr5-His6-Cys7, where the amino acids&#39; Aa1,2,6,7 positions are the potential metal-binding sites. Geometry-optimized stationary states of the ternary complexes and their products were selected from quantum chemistry calculations (presently the PM6 semi-empirical Hamiltonian) by comparing their electronic energies and their collision cross-sections (CCS) to those measured by ES-IM-MS. From the PM6 frequency calculations, the molecular parameters of the ternary complex and its products model the energy-dependent intensities of the two product channels using a competitive TCID method to determine the threshold energies of the reactions that relate to the 0 K enthalpies of dissociation (&#916;H0). Statistical mechanics thermal and entropy corrections using the PM6 rotational and vibrational frequencies provide the 298 K enthalpies of dissociation (&#916;H298). These methods describe an EI-IM-MS routine that can determine thermochemistry and equilibrium constants for a range of ternary metal ion complexes.

Thermochemical Studies of NiII and ZnII Ternary Complexes Using Ion Mobility-Mass Spectrometry

This article describes an experimental protocol using electrospray-ion mobility-mass spectrometry, semi-empirical quantum calculations, and energy-resolved threshold collision-induced dissociation to measure the relative thermochemistry of the dissociation of related ternary metal complexes.

A Study of the Complexation of Mercury(II) with Dicysteinyl Tetrapeptides by Electrospray Ionization Mass Spectrometry

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