SC thanks the Whitehead Summer Research Fellowship. MJS thanks the Startup Funds and the Faculty Development Fund at the University of San Francisco. MCH acknowledges funding from the National Institutes of Health (NIH MIRA 5R35GM133684-02) and the National Science Foundation (NSF CAREER 2048265).

1. Electronic absorption spectroscopy: direct titration with buffer competition
<ol>
	<li>Sample preparation
	<ol>
		<li>Prepare a buffered solution of 50 mM 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol (bisTris) at pH 7.4 using ultrapure water (&#62;18 M&#937; resistance). Remove trace metal ions by incubating with a high-affinity resin for at least 2 h with subsequent filtration.</li>
		<li>Dissolve or dilute a known quantity of the peptide into the metal-free buffer. 
		NOTE: When monitoring d-d bands with small extinction coefficients21, higher concentrations of peptide must be used. Here, the final concentration of C-peptide in buffered solution was 300 &#181;M (volume depends on the size of the cuvette). C-peptide was synthesized by solid-phase peptide synthesis and is detailed elsewhere in literature14.</li>
		<li>Dissolve a known mass of CuCl2 in ultrapure water to make a solution at 10-15 mM. 
		NOTE: It is important to initially dissolve the metal salt in nonbuffered water to prevent precipitation. Other Cu(II) salts may be used, but care must be taken to ensure that the anion is weakly coordinating.</li>
	</ol>
	</li>
	<li>Running the experiment
	<ol>
		<li>Turn on the electronic absorption spectrophotometer and let it warm up for ~15-20 min before use. Launch the spectrophotometer software and configure the parameters such as scanning range (200-900 nm), scan rate (200 nm/s), and double beam baseline-corrected (more parameters are listed in the Supplemental File).</li>
		<li>Collect a baseline with no cuvettes or samples in the beam paths.</li>
		<li>Using two matched cuvettes in the double-beam spectrophotometer, load one cuvette with 115 &#181;L of ultrapure water and the other cuvette with 115 &#181;L of the peptide sample. Ensure that there are no air bubbles in the cuvettes as these will interfere with the signal.</li>
		<li>Place the cuvette with ultrapure water in the reference beam and the cuvette with peptide in the sample beam.</li>
		<li>Collect the absorption spectrum of the metal-free (apo) peptide.</li>
		<li>Add in a sub-stoichiometric amount (0.5 equivalents, 150 &#181;M) of the Cu(II) solution into the cuvette with the peptide sample. Ensure that the volume of Cu(II) added is less than 3 &#181;L and record the volume for analysis later.</li>
		<li>Gently pipette up and down to mix the solution while avoiding the generation of air bubbles. Let the solution react and equilibrate for 5 min and record the absorption spectrum.</li>
		<li>Repeat the addition of Cu(II) aliquots as in step 1.2.6 into the peptide solution for the following equivalents: 1.0, 1.5, 2.0, 3.0, and 5.0 (or a total of 300, 450, 600, 900, and 1,500 &#181;M). Make sure to record the total volume of Cu(II) added and the total volume of the cuvette. 
		NOTE: If more resolution is desired, reduce the spacing between equivalents.</li>
		<li>Remove the sample cuvette and thoroughly clean as per the manufacturer&#39;s directions.</li>
		<li>Add the buffered solution without peptide and record the absorption spectrum. Repeat the addition of Cu(II) aliquots as in steps 1.2.5-1.2.8, recording the absorption spectrum for each Cu(II) equivalent.</li>
		<li>Export all spectra as csv files for processing. Thoroughly clean the cuvettes according to the manufacturer&#39;s instructions and power down the spectrophotometer.</li>
	</ol>
	</li>
	<li>Processing the data
	<ol>
		<li>Load all spectra on a spreadsheet program.</li>
		<li>Subtract the buffer-only (0 &#181;M Cu(II)) spectrum from every other spectrum to remove any absorbance features from the buffer itself.</li>
		<li>Normalize each spectrum to account for the dilution resulting from the addition of the Cu(II) solution (step 1.2.6). See the Supplemental File, Eq (1)14 for an example of the normalization where vinitial is the volume (115 &#181;L) of peptide added to the cuvette, vCu(II) is the volume of Cu(II) solution added in step 1.2.6, and Absbuffer subtracted spectrum is the data obtained in step 1.2.7.</li>
		<li>Graph all spectra together to identify regions of change. 
		&#8203;NOTE: Typical d-d bands from Cu(II) complexes range from 500 to 750 nm. This spectrophotometric titration can be challenging due to the small extinction coefficient from d-d bands, which are Laporte-forbidden transitions in octahedral geometry21. If the absorbance is too weak, an alternative approach is to utilize chromophoric ligands that result in charge transfer bands upon binding to Cu(II) (see section 2).</li>
	</ol>
	</li>
</ol>
2. Electronic absorption spectroscopy: peptide competition with chromophoric ligand
<ol>
	<li>Sample preparation
	<ol>
		<li>Dissolve 1,10-phenanthroline (phen) in ultrapure water to obtain a final concentration of ~1 mM.</li>
		<li>In addition to the sample preparation described in section 1.1 for the peptide (step 1.1.2) and Cu(II) (step 1.1.3), prepare a 10 &#181;M Cu(II) and 40 &#181;M phen solution in buffer ([Cu(phen)3]2+). Ensure that the volume fills the cuvette.</li>
	</ol>
	</li>
	<li>Running the experiment
	<ol>
		<li>Start the electronic absorption spectrophotometer as in steps 1.2.1 and 1.2.2 but set the scanning range to 200-400 nm.</li>
		<li>In two matched cuvettes, load one cuvette with 115 &#181;L of ultrapure water and the other cuvette with 115 &#181;L of the [Cu(phen)3]2+ solution. Place the cuvette with water in the reference beam and the cuvette with [Cu(phen)3]2+ solution in the sample beam.</li>
		<li>Collect the absorption spectrum of the metal-ligand complex.</li>
		<li>Add a stoichiometric amount (&#8776;1 equivalents, &#8776;10 &#181;M) of peptide to the [Cu(phen)3]2+ solution. Gently pipette up and down to thoroughly mix but be careful not to introduce air bubbles. Record the volume of peptide added for future analysis. 
		NOTE: Using cuvettes that hold 115 &#181;L, adding 3.83 &#181;L of 300 &#181;M peptide yields a final peptide concentration of 9.7 &#181;M.</li>
		<li>Incubate the solution for 5 min to reach equilibrium. Record the absorption spectrum. 
		NOTE: If the binding affinity of the metal-peptide and metal-ligand are similar, the concentration of peptide added will need to be in large excess. Be sure to account for the total volume of peptide added for normalization.</li>
		<li>Repeat the addition of peptide aliquots into the [Cu(phen)3]2+ solution for the following approximate equivalents: 2, 3, 4, 5, 6, 7, 8, 9, 10, 12, 14, 16, 18, 22, and 26. Record the volume of peptide added so that the diluted concentration can be determined.</li>
		<li>Remove the sample and clean the cuvette thoroughly according to the manufacturer&#39;s instructions. Collect a spectrum of the buffer. Collect a spectrum of 50 &#181;M peptide in the buffer.</li>
		<li>Export all data as csv files for processing and clean the cuvettes according to the manufacturer&#39;s instructions.</li>
	</ol>
	</li>
	<li>Processing the data
	<ol>
		<li>Load the spectra on a spreadsheet program and subtract the buffer spectrum from the other spectra.</li>
		<li>Normalize the spectra following Supplemental File, Eq (2)14.</li>
		<li>Using the extinction coefficient for [Cu(phen)3]2+ at 265 nm (&#949;max = 90,000 M-1cm-1)22, determine the concentration of [Cu(phen)3]2+ with each addition of the peptide.</li>
		<li>For each addition of the peptide, determine the concentration of the Cu(II)-peptide complex by subtracting the remaining concentration of [Cu(phen)3]2+, as determined in step 2.3.2, from the initial concentration of [Cu(phen)3]2+ (at 0 &#181;M peptide).</li>
		<li>Calculate the concentration of free phen ligand by the equation in Supplemental File, Eq (3)14.</li>
		<li>Calculate the concentration of free peptide using Supplemental File, Eq (4)14, where [peptide]stock represents the undiluted peptide titrated into the cuvette, V1 represents the volume of stock peptide added, V2 is the total volume of the cuvette, and [Cu2+-peptide] is determined in step 2.3.4.</li>
		<li>Calculate the experimental binding affinity (Kex) using Eq (5) in Supplemental File23.</li>
		<li>Relate the dissociation constant of Cu(II)-peptide to Kex by Eq (6)23 in Supplemental File, where Kd,Cu(II)-phen = 1.0 &#215; 10-9 (see 22). Determine the average and standard deviation from all the determined dissociation constants. 
		&#8203;NOTE: Under a 4:1 ratio of phen:Cu(II), [Cu(phen)3]2+, [Cu(phen)2]2+, and [Cu(phen)]2+ exist in the solution, and the peptide will chelate Cu(II) away from the species ([Cu(phen)]2+) with the weakest binding22.</li>
	</ol>
	</li>
</ol>
3. Isothermal titration calorimetry
<ol>
	<li>Sample preparation
	<ol>
		<li>Prepare a buffered solution of 15 mM 3-morpholinopropane-1-sulfonic acid (MOPS) at pH 7.4 using ultrapure water (&#62;18 M&#937; resistance). Remove trace metal ions by incubating with a high-affinity resin for at least 2 h with subsequent vacuum filtration through a bottle-top 0.45 &#181;m membrane.</li>
		<li>Dissolve a known mass of CuCl2 in ultrapure water to prepare a solution of &#8776;50-100 mM. Dilute this Cu(II) solution into buffer to obtain a 1.0 mL solution with final concentration of 1.4 mM Cu(II). Record the exact volume of the CuCl2 solution used.</li>
		<li>Dissolve or dilute the peptide solution in buffer to make 450 &#181;L of a 154 &#181;M peptide solution. Ensure that the same proportion of additional ultrapure water from step 3.1.2 is added to the peptide solution, which will reduce heat of dilution and increase signal-to-noise.</li>
		<li>After preparation of the samples, ensure that the solutions are at the same pH and, if needed, adjust accordingly.</li>
		<li>Optional step: Degas the solutions to minimize microbubbles loaded into the ITC.</li>
	</ol>
	</li>
	<li>Running the experiment
	<ol>
		<li>Turn on the ITC. Launch the ITC software to run the instrument. Wait for the first initialization, which will ask to rehome the buret; then, follow the instructions on the screen.</li>
		<li>Remove the cover from the reference cell. Remove any water from the reference cell and rinse three times with 450 &#181;L of degassed ultrapure water.</li>
		<li>Slowly draw up ultrapure water to the 450 &#181;L mark of a loading syringe, taking care not to introduce air bubbles into the syringe. Insert the loading syringe into the reference cell until it is &#8776;1 mm from the bottom, and slowly inject part of the solution until 150 &#181;L remains in the loading syringe. Move the loading syringe plunger quickly up and down by &#8776;25 &#181;L several times to dislodge any bubbles on the cell surface. Slowly inject until the plunger reaches the 100 &#181;L mark on the loading syringe, thereby dispensing a total of 350 &#181;L of ultrapure water into the reference cell, and replace the reference cell cover.</li>
		<li>Remove any residual solution from the sample cell and load with 450 &#181;L of 10 mM ethylenediaminetetraacetic acid (EDTA) using a loading syringe. Soak for 10 min to ensure that trace metal ions are removed because the EDTA will bind trace metals.</li>
		<li>Remove the EDTA solution (with bound trace metal ions) and thoroughly rinse the loading syringe with copious amounts of ultrapure water.</li>
		<li>Clean the ITC in accordance with the manufacturer&#39;s directions, rinsing the sample cell with ultrapure water.</li>
		<li>Condition the sample cell by rinsing with 450 &#181;L of buffer at least three times.</li>
		<li>Remove the buffer that is conditioning the sample cell. Load the peptide solution into the sample cell using the loading syringe (follow step 3.2.3).</li>
		<li>Rinse the titration syringe with 200 &#181;L of buffered solution. To do this, remove the plunger and use a micropipette to pipette the buffer through the hole at the top of the glass titration syringe, through the syringe, and out the needle below.</li>
		<li>Fully insert the plunger into the titration syringe.</li>
		<li>Dip the tip of the titration syringe needle into the metal solution and slowly pull the plunger up, causing the metal solution to fill the syringe and result&#160;in a void volume at the top of the glass part of the titration syringe. Remove most of the void volume by rotating the titration syringe parallel to the floor, remove the plunger, and slightly tilt the glass part toward the floor. Give the titration syringe a gentle shake so that the solution moves to the end of the glass part of the titration syringe and fills most of the void volume, but ensure that 2-3 &#181;L of void volume remains. While keeping the syringe parallel to the floor, reinsert the plunger.</li>
		<li>Hold the titration syringe upright, dip the tip of the needle back into the metal solution, and push the plunger down until air ceases to come out of the needle. Load the titration syringe by slowly pulling up the plunger to just above the 50 &#181;L mark while keeping the tip of the needle in the solution.</li>
		<li>Carefully insert the glass part of the titration syringe into the buret and screw until finger-tight. When a small amount of solution comes out of the titration syringe due to compression of the plunger, use a light-duty delicate wiper to carefully absorb the solution without touching the tip of the needle.</li>
		<li>Insert the buret with titration syringe into the sample cell and fasten it securely.</li>
		<li>Set up the parameters on the ITC software. Starting on Instrument Control, set the stirring rate (typical stirring rates range from 150 to 350 RPM) and the temperature at which the experiment will be conducted (typically 25 &#176;C). Enter the syringe and cell concentrations in millimolar units under Experiment Details.</li>
		<li>In the Experiment Method section, select Incremental Titration. Click on Setup and specify 20 injections of 2.5 &#181;L. If more resolution is required to observe the binding event, increase the number of injections and decrease the volume per injection. Input the time spacing between each injection so that it is long enough for the signal to equilibrate and return to baseline, typically 300 s.</li>
		<li>Click the run button to start the experiment and specify where the data are saved.</li>
		<li>Upon completion of the experiment, clean the sample cell and titration syringe.</li>
		<li>Run all experiments in at least triplicate to ensure precise data collection.</li>
		<li>Run a control experiment where the metal solution is titrated into the buffered solution (in the absence of peptide) to ensure the heat of dilution from the metal ion is small and that there are no unaccounted-for equilibria. If the heat of dilution is large, consider a different buffer system, if possible.</li>
	</ol>
	</li>
	<li>Processing the data
	<ol>
		<li>Launch the ITC analysis software and load the data file for analysis.</li>
		<li>Navigate to the Baseline tab and inspect the thermogram. Note any exogenous heat evolved or absorbed from air bubbles or other artifacts in the thermogram. Look for spikes that are not due to injection of the metal solution.</li>
		<li>Ensure that the baseline generated by the analysis software follows the portion of the data after injection and equilibration. If it deviates, use Baseline Pivot Points to adjust the baseline. Ensure that the Integration Regions include the peak generated from injection of the metal, but occlude any air bubbles or artifacts in the thermogram found in step 2. Subtract the baseline from the thermogram. 
		NOTE: This type of manipulation is recommended only after multiple thermograms are collected, so the experimenter knows what is real data and what is an artifact, as adjusting the baseline and Integration Regions may drastically affect the data.</li>
		<li>Navigate to the Modeling window to begin fitting the data where the analysis software will show the integrated and concentration-normalized data for each injection.</li>
		<li>Left-click on the datum of the first injection to remove it from the fitting algorithm. 
		NOTE: This is common since there will be minor mixing between the titrant and sample cell solution leading to inexact molar dispensing of the first injection.</li>
		<li>In the Models section, select Blank (constant) in the Style dropdown menu, which is based on the final injection enthalpy and will be subtracted from every data point, accounting for the heat of dilution. In addition, select Independent (or the best model for the system) in the second Style dropdown menu to fit the data. 
		NOTE: For standard 1:1 binding interactions, the most common model is Independent.</li>
		<li>Fit the data using the two models by pressing the green play button with a &#931;. 
		NOTE: Another software to process ITC data is SEDPHAT24.</li>
		<li>Account for all competing equilibria in post hoc analysis previously reported by Grossoehme et al.20.</li>
	</ol>
	</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coordinating+properties+of+the+amide+bond.+stability+and+structure+of+metal+ion+complexes+of+peptides+and+related+ligands.">Coordinating properties of the amide bond. stability and structure of metal ion complexes of peptides and related ligands.</a> Chemical Reviews. 82 (4), 385-426 (1982).</li><li>Liu, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metalloproteins+containing+cytochrome,+iron-sulfur,+or+copper+redox+centers.">Metalloproteins containing cytochrome, iron-sulfur, or copper redox centers.</a> Chemical Reviews. 114 (8), 4366 (2014).</li><li>Grossoehme, N. E., Spuches, A. M., Wilcox, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+isothermal+titration+calorimetry+in+bioinorganic+chemistry.">Application of isothermal titration calorimetry in bioinorganic chemistry.</a> Journal of Biological Inorganic Chemistry. 15 (8), 1183-1191 (2010).</li><li>Miessler, G. L., Fischer, P. J., Tarr, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Inorganic Chemistry. , (2014).</li><li>Walger, E., Marlin, N., Molton, F., Mortha, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Study+of+the+direct+red+81+dye/copper(II)-phenanthroline+system.">Study of the direct red 81 dye/copper(II)-phenanthroline system.</a> Molecules. 23 (2), 1-23 (2018).</li><li>Kocy&#322;a, A., Pomorski, A., Kr&#281;&#380;el, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molar+absorption+coefficients+and+stability+constants+of+Zincon+metal+complexes+for+determination+of+metal+ions+and+bioinorganic+applications.">Molar absorption coefficients and stability constants of Zincon metal complexes for determination of metal ions and bioinorganic applications.</a> Journal of Inorganic Biochemistry. 176, 53-65 (2017).</li><li>Zhao, H., Piszczek, G., Schuck, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SEDPHAT+-+A+platform+for+global+ITC+analysis+and+global+multi-method+analysis+of+molecular+interactions.">SEDPHAT - A platform for global ITC analysis and global multi-method analysis of molecular interactions.</a> Methods. 76, 137-148 (2015).</li><li>NIST. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Critically Selected Stability Constants of Metal Complexes, Version 8.0. , (2004).</li><li>Riener, C. K., Kada, G., Gruber, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quick+measurement+of+protein+sulfhydryls+with+Ellman's+reagent+and+with+4,4&#8242;-dithiodipyridine.">Quick measurement of protein sulfhydryls with Ellman's reagent and with 4,4&#8242;-dithiodipyridine.</a> Analytical and Bioanalytical Chemistry. 373 (4-5), 266-276 (2002).</li><li>Bradford, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+and+sensitive+method+for+the+quantitation+of+microgram+quantities+of+protein+utilizing+the+principle+of+protein-dye+binding.">A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding.</a> Analytical Biochemistry. 72 (1-2), 248-254 (1976).</li></ol>

The authors declare no competing interests.

This article provides a robust method for quantifying the affinity and thermodynamics of Cu(II) binding to peptides. Complexes with Cu(II) are ideally suited to monitor the d-d absorption band at the metal site due to its d9 electron configuration. Although the extinction coefficient is small, thus requiring larger concentrations of the complex to yield a reliable signal, titrations of Cu(II) into peptide can quickly provide insight into the binding stoichiometry and approximate binding affinity. However, it c...

Biology has evolved to utilize the diverse chemistry of metal ions needed for life to adapt and survive in its surrounding environment. An estimated 25%-50% of proteins use metal ions for structure and function1. The particular role and redox state of the metal ion is directly related to the composition and geometry of the biological ligands that coordinate it. In addition, redox-active metal ions such as Cu(II) must be tightly regulated lest they interact with oxidizing agents via Fenton-like chemistry to form reactive oxygen species (ROS)2,3,4. Understanding the binding modes and affinity that drive its biochemistry should help elucidate the biological role of the metal ion.
Many techniques are used to study the binding interactions of metals and peptides. These are mostly spectroscopic techniques but also include computer simulations using molecular dynamics, as seen through Cu(II) interactions with a fragment of amyloid beta (A&#946;)5. A widely used spectroscopic technique that is accessible to many universities is nuclear magnetic resonance (NMR). By using the paramagnetic nature of Cu(II), Gaggelli et al. were able to show where the metal ion binds on a petide through relaxation of nearby nuclei6. Electron paramagnetic resonance (EPR) may also be utilized to probe the location and mode of the paramagnetic metal ion binding7. Other spectroscopic techniques such as circular dichroism (CD) can describe the coordination about Cu(II) in systems such as tripeptide systems8, and mass spectrometry can show stoichiometry and to which residues the metal ion is coordinated through fragmentation patterns9,10.
Some of these techniques, such as NMR, are label-free but require large concentrations of peptide, posing challenges for study. Another common technique called fluorescence spectroscopy has been utilized to relate the position of a tyrosine or tryptophan with quenching from a Cu(II)11,12. Similarly, this technique can show structural changes as a result of Cu(II) binding13. However, challenges with these metal-peptide binding studies are that they probe chromophoric amino acids such as tyrosine which not all systems have, that the metal ion binds under a classical model, and that the technique may not be conducive under physiological conditions. Indeed, several peptides are emerging that do not contain such chromophoric amino acids or bind under classical models, precluding the use of these techniques14,15. This article details approaches for assessing binding properties in these scenarios under physiologically relevant conditions.
Biological ligands may adopt different protonation states that can affect metal ion binding such as the imidazole ring on histidine. If pH is not consistently maintained, the results can be convoluted or conflicting. For this reason, buffers are an essential component in the study of metal-protein/peptide interactions. However, many buffers have been shown to favorably interact with metal ions16,17. In addition to competing with the biological molecule of interest, the buffer may have similar coordinating atoms that may be difficult to distinguish from the coordinating atoms of the peptide or protein. In this study, the focus is on electronic absorption spectroscopy and isothermal titration calorimetry (ITC) as two complementary techniques for studying Cu(II)-peptide interactions, with special considerations concerning buffer choice.
Electronic absorption spectroscopy is a rapid, widely accessible technique for studying metal-binding interactions. Irradiation with light in the ultraviolet (UV) or visible wavelengths can lead to absorption of metal-centered d-d bands, which provide valuable information on ligand classification, metal geometries, and apparent binding affinities18,19. For these complexes, direct titrations of metal ions into protein or peptide solutions can quantify binding stoichiometries and apparent binding affinities. In some cases, such as d5 or d10 electron configurations, the complex does not absorb light (i.e., is spectroscopically silent). In these spectroscopically silent transition metal complexes, these limitations can be circumvented by using a competing ligand that, upon coordinating to the metal ion, yields detectable charge transfer bands. In either case, this approach is limited to quantifying only stoichiometry and apparent binding affinity, and no insight into binding enthalpy is provided without approximations.
Complementing information obtained from electronic absorption spectroscopy, ITC is an attractive technique for direct and rigorous quantification of the binding enthalpy20. ITC directly measures the heat released or consumed during a binding event and, since the titration takes place at constant pressure, the heat measured is the enthalpy of all equilibria (&#916;HITC). In addition, the stoichiometry of the binding event (n) and the apparent binding affinity (KITC) are quantified. From these parameters, the free energy (&#916;GITC) and entropy (&#916;SITC) are determined, providing a thermodynamic snapshot of the binding event. As it does not rely on light absorption, ITC is an ideal technique for spectroscopically silent species, for example, d5 or d10 metal ion complexes. However, since calorimetry measures heat, any unmatched buffer systems and unaccounted-for equilibria may adversely affect the analysis to accurately determine the metal ion binding thermodynamics, and great care must be taken to address these factors20. If performed with the appropriate rigor, ITC is a robust technique for determining the thermodynamics of metal-protein/peptide complexes.
Here, a chromophorically silent copper-binding peptide, C-peptide, is used to demonstrate the complementary use of the two techniques. C-peptide is a 31 residue cleavage product (EAEDLQVGQVELGGGPGAGSLQPLALEGSLQ) formed during insulin maturation; it lacks chromophoric residues but has been shown to bind Cu(II) with physiologically relevant affinity14,15. The Cu(II) binding site consists of the side chains of a glutamate and an aspartate as well as the N-terminus of the peptide14,15. These coordinating atoms closely resemble those of many commonly used buffered systems. Here, the tandem use of the d-d and charge transfer bands in electronic absorption spectroscopy and ITC in quantifying the Cu(II) binding thermodynamics to C-peptide is shown. The approach from studying Cu(II) binding to C-peptide can be applied to other metal ions and protein/peptide systems.

<table><tbody><tr><td>1,10-phenanthroline</td><td>Sigma Aldrich</td><td>131377-25G</td><td></td></tr><tr><td>bis-Tris buffer</td><td>Fisher</td><td>BP301-100</td><td></td></tr><tr><td>Bottle-top 0.45 micron membrane</td><td>Nalgene</td><td>296-4545</td><td>Any filtration system that removes the resin without introducing contaminants is acceptable</td></tr><tr><td>Copper(II) chloride</td><td>Alfa Aesar</td><td>12458</td><td></td></tr><tr><td>EDTA</td><td>Sigma Aldrich</td><td>EDS-500G</td><td></td></tr><tr><td>Electronic absorption spectrophotometer</td><td>Varian</td><td>Cary 5000</td><td>Another suitable sensitive spectrophotometer is acceptable</td></tr><tr><td>high affinity resin</td><td>Sigma Aldrich</td><td>C7901-25G</td><td></td></tr><tr><td>Isothermal titration calorimeter (ITC)</td><td>TA Instruments</td><td>Nano ITC Low Volume</td><td></td></tr><tr><td>ITC analysis software</td><td>TA Instruments</td><td>NanoAnalyze</td><td>SEDPHAT (&lt;em&gt;Methods&lt;/em&gt;. &lt;strong&gt;2015&lt;/strong&gt;, 76: 137&amp;ndash;148) may also be used</td></tr><tr><td>ITC software</td><td>TA Instruments</td><td>ITCRun</td><td></td></tr><tr><td>light-duty delicate wiper</td><td>Kimwipe</td><td>34155</td><td></td></tr><tr><td>loading syringe</td><td>Hamilton</td><td>Syr 500 uL, 1750 TLL-SAL</td><td></td></tr><tr><td>matched cuvettes</td><td>Starna Cells, Inc</td><td>16.100-Q-10/Z20</td><td>Ensure that the window for the small volume cuvette matches the beam height of the spectrophotometer</td></tr><tr><td>MOPS buffer</td><td>Alfa Aesar</td><td>A12914</td><td></td></tr><tr><td>spectrophotometer software</td><td>Cary</td><td>WinUV Scan</td><td></td></tr><tr><td>spreadsheet program</td><td>Microsoft</td><td>Excel</td><td>Any suitable spreadsheet program will work</td></tr><tr><td>titration syringe</td><td>TA Instruments</td><td>5346</td><td></td></tr><tr><td>ultrapure water</td><td>Millipore Sigma</td><td>Milli-Q</td><td>Any water is okay as long as &gt;18 M&amp;Omega; resistance</td></tr></tbody></table>

quantifying the binding interactions between cu(ii) and peptide residues in the presence and absence of chromophores

Copper(II) is an essential metal in biological systems, conferring unique chemical properties to the biomolecules with which it interacts. It has been reported to directly bind to a variety of peptides and play both necessary and pathological roles ranging from mediating structure to electron transfer properties to imparting catalytic function. Quantifying the binding affinity and thermodynamics of these Cu(II)-peptide complexes in vitro provides insight into the thermodynamic driving force of binding, potential competitions between different metal ions for the peptide or between different peptides for Cu(II), and the prevalence of the Cu(II)-peptide complex in vivo. However, quantifying the binding thermodynamics can be challenging due to a myriad of factors, including accounting for all competing equilibria within a titration experiment, especially in cases where there are a lack of discrete spectroscopic handles representing the peptide, the d-block metal ion, and their interactions.
Here, a robust set of experiments is provided for the accurate quantification of Cu(II)-peptide thermodynamics. This article focuses on the use of electronic absorption spectroscopy in the presence and absence of chromophoric ligands to provide the needed spectroscopic handle on Cu(II) and the use of label-free isothermal titration calorimetry. In both experimental techniques, a process is described to account for all competing equilibria. While the focus of this article is on Cu(II), the described set of experiments can apply beyond Cu(II)-peptide interactions, and provide a framework for accurate quantification of other metal-peptide systems under physiologically relevant conditions.

The goal was to quantify and corroborate the thermodynamics of Cu(II) binding to C-peptide using the complementary techniques of electronic absorption spectroscopy and ITC. Due to the robust nature of electronic absorption spectroscopy, a direct titration of Cu(II) into 300 &#181;M C-peptide was performed (Figure 1). Addition of 150 &#181;M of Cu(II) caused an immediate increase in the band at 600 nm, attributed to the d-d band of Cu(II), and continued to increase until 300 &#181;M Cu(II) wa...

Watch this Scientific Journal Video about Quantifying the Binding Interactions Between CuII and Peptide Residues in the Presence and Absence of Chromophores at JoVE.com

Quantifying the Binding Interactions Between CuII and Peptide Residues in the Presence and Absence of Chromophores

This article focuses on the use of electronic absorption spectroscopy and isothermal titration calorimetry to probe and quantify the thermodynamics of Cu(II) binding to peptides and proteins.

Quantifying the Binding Interactions Between Cu(II) and Peptide Residues in the Presence and Absence of Chromophores

Copper(II) is an essential metal in biological systems, conferring unique chemical properties to the biomolecules with which it interacts. It has been ...

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Research

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Biochemistry

2.3K Views.  University of San Francisco. This article focuses on the use of electronic absorption spectroscopy and isothermal titration calorimetry to probe and quantify the thermodynamics of Cu(II) binding to peptides and proteins.

Video: Quantifying the Binding Interactions Between CuII and Peptide Residues in the Presence and Absence of Chromophores

Identification of Small Molecule-binding Proteins in a Native Cellular Environment by Live-cell Photoaffinity Labeling

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several target identification methods have been developed and used to successfully elucidate the binding proteins of a variety of small molecules, these techniques have drawbacks that make them unsuitable for detecting certain types of small molecule-target interactions. In particular, non-covalent interactions that depend on native cellular conditions, such as those of membrane proteins whose structures may be perturbed upon cell lysis, are often not amenable to affinity-based target identification methods. Here, we demonstrate a method wherein a probe containing a photolabile group is used to covalently crosslink to the small molecule binding protein within the environment of the live cell, allowing the detection and isolation of the target protein without the need for maintenance of the interaction after cell lysis. This technique is a valuable tool for studying biologically interesting small molecules with unknown mechanisms, both in the context of basic biology as well as drug discovery.

We describe here a method for identification of small molecule-binding proteins using photoaffinity labeling. The advantage of this technique is that binding and covalent labeling of the target proteins occurs within the live cellular environment, removing the risk of disrupting native protein structure and binding conditions upon cell lysis.

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several ...

Method for Identifying Small Molecule Inhibitors of the Protein-protein Interaction Between HCN1 and TRIP8b

Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels are expressed ubiquitously throughout the brain, where they function to regulate the excitability of neurons. The subcellular distribution of these channels in pyramidal neurons of hippocampal area CA1 is regulated by tetratricopeptide repeat-containing Rab8b interacting protein (TRIP8b), an auxiliary subunit. Genetic knockout of HCN pore forming subunits or TRIP8b, both lead to an increase in antidepressant-like behavior, suggesting that limiting the function of HCN channels may be useful as a treatment for Major Depressive Disorder (MDD). Despite significant therapeutic interest, HCN channels are also expressed in the heart, where they regulate rhythmicity. To circumvent off-target issues associated with blocking cardiac HCN channels, our lab has recently proposed targeting the protein-protein interaction between HCN and TRIP8b in order to specifically disrupt HCN channel function in the brain. TRIP8b binds to HCN pore forming subunits at two distinct interaction sites, although here the focus is on the interaction between the tetratricopeptide repeat (TPR) domains of TRIP8b and the C terminal tail of HCN1. In this protocol, an expanded description of a method for purifying TRIP8b and executing a high throughput screen to identify small molecule inhibitors of the interaction between HCN and TRIP8b, is described. The method for high throughput screening utilizes a Fluorescence Polarization (FP) -based assay to monitor the binding of a large TRIP8b fragment to a fluorophore-tagged eleven amino acid peptide corresponding to the HCN1 C terminal tail. This method allows &#39;hit&#39; compounds to be identified based on the change in the polarization of emitted light. Validation assays are then performed to ensure that &#39;hit&#39; compounds are not artifactual.

The interaction between HCN channels and their auxiliary subunit has been identified as a therapeutic target in Major Depressive Disorder. Here, a fluorescence polarization-based method for identifying small molecule inhibitors of this protein-protein interaction, is presented.

Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels are expressed ubiquitously throughout the brain, where they function to regulate the ...

Structure and Coordination Determination of Peptide-metal Complexes Using 1D and 2D 1H NMR

Copper (I) binding by metallochaperone transport proteins prevents copper oxidation and release of the toxic ions that may participate in harmful redox reactions. The Cu (I) complex of the peptide model of a Cu (I) binding metallochaperone protein, which includes the sequence MTCSGCSRPG (underlined is conserved), was determined in solution under inert conditions by NMR spectroscopy.
NMR is a widely accepted technique for the determination of solution structures of proteins and peptides. Due to difficulty in crystallization to provide single crystals suitable for X-ray crystallography, the NMR technique is extremely valuable, especially as it provides information on the solution state rather than the solid state. Herein we describe all steps that are required for full three-dimensional structure determinations by NMR. The protocol includes sample preparation in an NMR tube, 1D and 2D data collection and processing, peak assignment and integration, molecular mechanics calculations, and structure analysis. Importantly, the analysis was first conducted without any preset metal-ligand bonds, to assure a reliable structure determination in an unbiased manner.

Structure and Coordination Determination of Peptide-metal Complexes Using 1D and 2D 1H NMR

The NMR-solution structure of a metallochaperone model peptide with Cu (I) was determined, and a detailed protocol from sample preparation and 1D and 2D data collection to a three-dimensional structure is described.

Copper (I) binding by metallochaperone transport proteins prevents copper oxidation and release of the toxic ions that may participate in harmful redox ...

Chemistry

Analysis of AtHIRD11 Intrinsic Disorder and Binding Towards Metal Ions by Capillary Gel Electrophoresis and Affinity Capillary Electrophoresis

Plants are strongly dependent on their environment. In order to adjust to stressful changes (e.g., drought and high salinity), higher plants evolve classes of intrinsically disordered proteins (IDPs) to reduce oxidative and osmotic stress. This article uses a combination of capillary gel electrophoresis (CGE) and mobility shift affinity electrophoresis (ACE) in order to describe the binding behavior of different conformers of the IDP AtHIRD11 from Arabidopsis thaliana. CGE is used to confirm the purity of AtHIRD11 and to exclude fragments, posttranslational modifications, and other impurities as reasons for complex peak patterns. In this part of the experiment, the different sample components are separated by a viscous gel inside a capillary by their different masses and detected with a diode array detector. Afterward, the binding behavior of the sample towards various metal ions is investigated by ACE. In this case, the ligand is added to the buffer solution and the shift in migration time is measured in order to determine whether a binding event has occurred or not. One of the advantages of using the combination of CGE and ACE to determine the binding behavior of an IDP is the possibility to automate the gel electrophoresis and the binding assay. Furthermore, CGE shows a lower limit of detection than the classical gel electrophoresis and ACE is able to determine the manner of binding a ligand in a fast manner. In addition, ACE can also be applied to other charged species than metal ions. However, the use of this method for binding experiments is limited in its ability to determine the number of binding sites. Nevertheless, the combination of CGE and ACE can be adapted for characterizing the binding behavior of any protein sample towards numerous charged ligands.

This protocol combines the characterization of a protein sample by capillary gel electrophoresis and a fast-binding screening for charged ligands by affinity capillary electrophoresis. It is recommended for proteins with a flexible structure, such as intrinsically disordered proteins, to determine any differences in binding for different conformers.

Plants are strongly dependent on their environment. In order to adjust to stressful changes (e.g., drought and high salinity), higher plants evolve ...

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen Presenting Cells (APCs). This is despite T-cell antigen receptors (TCRs) exerting usually a moderate affinity (&#181;M range) to antigen when binding is measured in vitro1. In view of the molecular and cellular parameters contributing to T-cell antigen sensitivity, a microscopy-based methodology has been developed as a means to monitor TCR-pMHC binding in situ, as it occurs within the synapse of a live T-cell and an artificial and functionalized glass-supported planar lipid bilayer (SLB), which mimics the cell membrane of an Antigen presenting Cell (APC) 2. Measurements are based on F&#246;rster Resonance Energy Transfer (FRET) between a blue- and red-shifted fluorescent dye attached to the TCR and the pMHC. Because the efficiency of FRET is inversely proportional to the sixth power of the inter-dye distance, one can employ FRET signals to visualize synaptic TCR-pMHC binding. The sensitive of the microscopy approach supports detection of single molecule FRET events. This allows to determine the affinity and off-rate of synaptic TCR-pMHC interactions and in turn to interpolate the on-rate of binding. Analogous assays could be applied to measure other receptor-ligand interactions in their native environment.

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

This manuscript describes how to conduct (single molecule) F&#246;rster Resonance Energy Transfer (FRET)- based assays to measure the binding dynamics between T-cell antigen receptor (TCR) and antigenic peptide-loaded MHC molecules as they occur within the immunological synapse of a T-cell in contact with a functionalized planar supported lipid bilayer.

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen ...

Bioengineering

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, ...

Covalent Labeling with Diethylpyrocarbonate for Studying Protein Higher-Order Structure by Mass Spectrometry

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool for this purpose, especially for protein systems that are difficult to study by traditional methods. To study a protein&#39;s structure by MS, specific chemical reactions are performed in solution that encode a protein&#39;s structural information into its mass. One particularly effective approach is to use reagents that covalently modify solvent accessible amino acid side chains. These reactions lead to mass increases that can be localized with residue-level resolution when combined with proteolytic digestion and tandem mass spectrometry. Here, we describe the protocols associated with use of diethylpyrocarbonate (DEPC) as a covalent labeling reagent together with MS detection. DEPC is a highly electrophilic molecule capable of labeling up to 30% of the residues in the average protein, thereby providing excellent structural resolution. DEPC has been successfully used together with MS to obtain structural information for small single-domain proteins, such as &#946;2-microglobulin, to large multi-domain proteins, such as monoclonal antibodies.

The experimental procedures for performing diethylpyrocarbonate-based covalent labeling with mass spectrometric detection are described. Diethylpyrocarbonate is simply mixed with the protein or protein complex of interest, leading to the modification of solvent accessible amino acid residues. The modified residues can be identified after proteolytic digestion and liquid chromatography/mass spectrometry analysis.

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool ...

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.

This article describes an experimental protocol using electrospray-ion mobility-mass spectrometry (ES-IM-MS) and energy-resolved threshold ...

Optimized Methods for the Surface Immobilization of Collagens and Collagen Binding Assays

Fibrosis occurs in various tissues as a reparative response to injury or damage. If excessive, however, fibrosis can lead to tissue scarring and organ failure, which is associated with high morbidity and mortality. Collagen is a key driver of fibrosis, with type I and type III collagen being the primary types involved in many fibrotic diseases. Unlike conventional protocols used to immobilize other proteins (e.g., elastin, albumin, fibronectin, etc.), comprehensive protocols to reproducibly immobilize different types of collagens in order to produce stable coatings are not readily available. Immobilizing collagen is surprisingly challenging because multiple experimental conditions may affect the efficiency of immobilization, including the type of collagen, the pH, the temperature, and the type of microplate used. Here, a detailed protocol to reproducibly immobilize and quantify type I and III collagens resulting in&#160;stable and reproducible gels/films is provided. Furthermore, this work demonstrates how to perform, analyze, and interpret in vitro time-resolved fluorescence binding studies to investigate the interactions between collagens and candidate collagen-binding compounds (e.g., a peptide conjugated to a metal chelate carrying, for example, europium [Eu(III)]). Such an approach can be universally applied to various biomedical applications, including&#160;the field of molecular imaging to develop targeted imaging probes, drug development, cell toxicity studies, cell proliferation studies, and immunoassays.

This work presents an optimized protocol to reproducibly immobilize and quantify type I and III collagen onto microplates, followed by an improved in vitro binding assay protocol to study collagen-compound interactions using a time-resolved fluorescence method. The subsequent step-by-step data analysis and data interpretation are provided.

Fibrosis occurs in various tissues as a reparative response to injury or damage. If excessive, however, fibrosis can lead to tissue scarring and organ ...

A High Throughput MHC II Binding Assay for Quantitative Analysis of Peptide Epitopes

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or design1,2. Here, a peptide-MHC II binding assay is scaled to 384-well format. The scaled down protocol reduces reagent costs by 75% and is higher throughput than previously described 96-well protocols1,3-5. Specifically, the experimental design permits robust and reproducible analysis of up to 15 peptides against one MHC II allele per 384-well ELISA plate. Using a single liquid handling robot, this method allows one researcher to analyze approximately ninety test peptides in triplicate over a range of eight concentrations and four MHC II allele types in less than 48 hr. Others working in the fields of protein deimmunization or vaccine design and development may find the protocol to be useful in facilitating their own work. In particular, the step-by-step instructions and the visual format of JoVE should allow other users to quickly and easily establish this methodology in their own labs.

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or design. &#160;Here, a peptide-MHC II binding assay scaled to 384-well plates is described. This cost effective format should prove useful in the fields of protein deimmunization and vaccine design and development.

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or ...

Quantifying the Binding Interactions Between Cu(II) and Peptide Residues in the Presence and Absence of Chromophores

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A High Throughput MHC II Binding Assay for Quantitative Analysis of Peptide Epitopes