Investigation into the Molecular and Thermodynamic Basis of

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Investigation into the Molecular and Thermodynamic Basis of Protein Interactions in Multimodal Chromatography Using Functionalized Nanoparticles Kartik Srinivasan,†,‡ Siddharth Parimal,†,‡ Maria M. Lopez,‡ Scott A. McCallum,‡,§ and Steven M. Cramer*,†,‡ †

Howard P. Isermann Department of Chemical and Biological Engineering, ‡Center for Biotechnology and Interdisciplinary Studies, and §Department of Biology, Rensselaer Polytechnic Institute, Troy, New York 12180, United States S Supporting Information *

ABSTRACT: Although multimodal chromatography offers significant potential for bioseparations, there is a lack of molecular level understanding of the nature of protein binding in these systems. In this study a nanoparticle system is employed that can simulate a chromatographic resin surface while also being amenable to isothermal titration calorimetry (ITC) and solution NMR. ITC and NMR titration experiments are carried out with 15N-labeled ubiquitin to investigate the interactions of ubiquitin with nanoparticles functionalized with two industrially important multimodal ligands. The ITC results suggest that binding to both multimodal ligand surfaces is entropically driven over a range of temperatures and that this is due primarily to the release of surface bound waters. In order to reveal structural details of the interaction process, bindinginduced chemical shift changes obtained from the NMR experiments are employed to obtain dissociation constants of individual amino acid residues on the protein surface. The residue level information obtained from NMR is then used to identify a preferred binding face on ubiquitin for interaction to both multimodal ligand surfaces. In addition, electrostatic potential and spatial aggregation propensity maps are used to determine important protein surface property data that are shown to correlate well with the molecular level information obtained from NMR. Importantly, the data demonstrate that the cluster of interacting residues on the protein surface act co-operatively to give rise to multimodal binding affinities several orders of magnitude greater than those obtained previously for interactions with free solution ligands. The use of NMR and ITC to study protein interactions with functionalized nanoparticles offers a new tool for obtaining important molecular and thermodynamic insights into protein affinity in multimodal chromatographic systems.

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INTRODUCTION Production of high-purity biotherapeutics requires the development of efficient bioseparation processes. The creation of highresolution separations for complex bioanalytical purposes is also of great interest. The majority of chromatographic media operate by a single interaction mode, with common chromatographic resins employing either electrostatic (e.g., ion exchange) or hydrophobic interactions (e.g., hydrophobic interaction, reversed phase). However, recent advances in the design of multimodal (or mixed mode) chromatographic systems have produced new materials that create unique windows of selectivity as compared to traditional single mode chromatographic systems by offering a combination of different interaction types within a single resin.1−7 Importantly, these new classes of resins have been shown to improve product quality and process efficiency in industrial-scale manufacturing.8,9 Analytical scale multimodal (MM) systems have also shown utility for complex analytical separations when coupled to mass spectrometry.10,11 © XXXX American Chemical Society

The majority of the multimodal (or mixed-mode) resins that are commercially available employ various combinations of hydrophobic, electrostatic, and/or hydrogen bonding interactions. A wide variety of moieties and geometries are currently being explored, and the combinatorial effects from mixing different interaction modes can create unexpected selectivity trends. The complex nature of the interactions involved between these ligands and proteins has hindered the optimal use of these materials and the design of improved multimodal ligands. Accordingly, there is an urgent need to develop both experimental and simulation methods to provide a deeper understanding of the interactions and selectivity in these systems.12,13 Developments in NMR spectroscopy, combined with the use of stable isotopes (13C, 15N, and 1H), have facilitated the use of Received: January 31, 2014 Revised: October 10, 2014

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dx.doi.org/10.1021/la502141q | Langmuir XXXX, XXX, XXX−XXX

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Figure 1. (A) Schematic representation of the nanoparticle structure and the protein−NP surface interaction. Particle surface chemistry of (B) “Capto ligand” and (C) “Nuvia ligand”.

the ligands as well as avidity effects which can play an important role in solid phase chromatographic systems. Hence, in order to examine the effects of co-operativity associated with the interacting residues at the binding interface, it was essential to probe the nature of protein interactions with solid ligand surfaces. The binding of proteins to solid surfaces has been studied using a range of experimental techniques including infrared spectroscopy, circular dichroism (CD), fluorescence, and NMR.21 Solid state NMR has been employed to determine the structure of the terminal helix of statherin when bound to hydroxyapatite.22 A combination of 1H NMR and CD was used to study the adsorption of a model 13 residue peptide to charged substrates.23 Atomic force microscopy (AFM) has also been employed to examine the behavior of ferritin at solid interfaces.24 Moreover, NMR CSPs have been employed to identify the ubiquitin−gold nanoparticle interaction site.25 Further, hydrogen−deuterium exchange (HDX) mass spectroscopy has been used to identify solvent accessible regions on proteins during binding to chromatographic surfaces.26,27 HDX has also been used in concert with 2D NMR spectroscopy to

a variety of NMR-based techniques for investigating protein− ligand interactions with atomic resolution.14,15 The use of ligand-induced chemical shift perturbation (CSP) to monitor protein−ligand interactions is a well-established technique in NMR spectroscopy.16,17 CSPs are extremely sensitive to local electronic environment effects, making it possible to detect ligand complexation at the atomic level while also providing an accurate determination of binding affinity.18,19 We have examined ubiquitin binding in ion exchange and multimodal chromatographic systems using solution phase NMR experiments to examine the interactions of ubiquitin and model untethered multimodal ligands as well as chromatographic data for a library of ubiquitin mutants.12 In addition, we have used molecular dynamics (MD) simulations to provide additional insights into the interactions of MM ligands with ubiquitin.13,20 This previous work has identified multiple binding residues in close proximity on the surface of ubiquitin which has led to the identification of a preferred binding face for the interaction of this protein with MM ligands. Although residue level information was obtained with these previous studies, these approaches did not account for the effects of immobilization of B


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Scheme 1

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study the conformational changes associated with protein adsorption to polystyrene nanospheres.28 NMR spectroscopy has been employed in concert with CD to probe changes in protein conformation on adsorption to silica nanoparticles of different diameters.29 Finally, several investigators have also examined the thermodynamic driving forces associated with the adsorption of proteins to solid surfaces such as silica and polystyrene particles and have studied the effect of adsorption on the structure and stability of proteins through the use of CD and differential scanning calorimetry (DSC).30−32 Self-assembled monolayers (SAMs) have emerged as an important platform to probe protein−surface interactions.33−36 SAMs enable the precise control of the chemical properties of a solid surface by varying the concentrations and relative compositions of the functionalized head groups.36 SAMs have also been used in concert with nanoparticles (NPs) to provide useful systems for modeling solid surfaces.37−40 Amino acidfunctionalized gold NPs have been shown to be an effective platform for studying protein−surface interactions.39,40 In the present study, a pseudo-solid-state resin system is created using gold NPs functionalized with SAMs containing MM ligands (Figure 1A). To investigate the thermodynamics of protein interactions with MM surfaces and to probe these binding processes at the molecular level, isothermal titration calorimetry (ITC) and NMR spectroscopy are employed. The NP system thus employed not only enables us to mimic MM chromatographic resin surfaces but also provides an experimental approach that is amenable to ITC and solution NMR. ITC experiments are performed to examine the thermodynamic driving forces involved in the interaction of human ubiquitin with the MM ligand functionalized nanoparticles (MM NPs). Further, NMR titration experiments are carried out using 15Nenriched human ubiquitin with the multimodal nanoparticles (MM NP) to determine the primary sites of specific protein− ligand interactions and the dissociation constants for the amino acid residues at the binding interface. Finally, electrostatic potential (EP) and spatial aggregation propensity (SAP) maps are generated to compare protein surface property data with the residue level binding information obtained from NMR. The use of solution NMR in concert with ITC on these pseudo-solidstate systems represents a new approach for studying the structural and thermodynamic basis of protein adsorption in MM chromatography.

EXPERIMENTAL SECTION

Materials. Sephadex G-50 Superfine media was packed into a BioRad glass chromatography column (5 cm × 16 cm). A SP Sepharose FF HiLoad16/10 column was obtained from Pharmacia (GE Healthcare). A Bondapak C18 Prep column (7.8 mm × 300 mm) was obtained from Waters. Stable isotopically labeled ammonium chloride was purchased from Isotec (Sigma-Aldrich). Hexaethylene glycol thiols terminated with N-hydroxysuccinimide ester groups were purchased from Prochimia (Poland). Acetonitrile was purchased from Acros Organics. Iron(III) chloride, 4-aminohippuric acid, ammonium chloride, potassium phosphate, potassium sulfate, calcium chloride, tris(hydroxymethyl)aminomethane, hydrochloric acid, protease cocktail inhibitor, deoxyribonuclease (DNAse), sodium chloride, sodium azide, acetic acid, sodium acetate, deuterium oxide (D2O), 3(trimethylsilyl)propionic acid-d4 sodium salt (TMSP), and NMR tubes were purchased from Sigma-Aldrich. Magnesium chloride (hexahydrate) was purchased from Mallinckrodt Baker. Ampicillin and IPTG was obtained from Gold Biotechnology. MOPS, tricine, dextrose, thiamine hydrochloride, sodium hydroxide, and sterile Nalgene filter packs were purchased from Thermo Fisher Scientific. Centriprep centrifugal filter devices were purchased from Millipore. Nbenzoyllysine was purchased from Chem-Impex International. Equipment. Purification of ubiquitin was carried using an Ä kta Explorer 100 equipped with Unicorn Software 3.1 from GE Healthcare. Reverse phase chromatography was performed using a Waters high performance liquid chromatography system consisting of a 600 multisolvent delivery system, a 717 Waters Intelligent Sample Processor autoinjector, and a 996 photodiode array detector controlled by a Millennium chromatography software manager. Synthesis of MM Ligand Functionalized Nanoparticles. The MM ligand head groups, N-benzoyllysine (2) and 4-aminohippuric acid (3), were employed to create the “Capto ligand” 4 and “Nuvia ligand” 5 linkers, respectively (Scheme 1). The activated Nhydroxysuccinimide ester (1) was dissolved in THF. The MM ligand headgroup was dissolved in PBS, pH 7.5, at a 20 M excess concentration to the activated ester. The NHS ester solution was then added to the ligand solution, and the mixture was stirred at room temperature for 36 h. The final product was purified from the excess unreacted ligands using reverse phase liquid chromatography operated with an acetonitrile gradient as described below. The purified MM ligand thiol was further utilized to functionalize the gold NPs. The MM ligand functionalized gold NPs were prepared by a placeexchange process of the synthesized MM ligand thiols with 1pentanethiol functionalized gold NPs (diameter ∼2.5 nm, Figures S1 and S2, Supporting Information) produced according to the published procedure.40−43 Quantitative proton NMR (using TMSP as a calibrated standard) was carried out to confirm the functionalization of the NPs and to calculate the surface density of the ligand head C


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was also carried for pure ubiquitin at a concentration of 200 μM in the same buffer. The resulting scattering data were fit (Dynamics software; Wyatt) by assuming the species in solution to be Rayleigh spheres. Protein Surface Characterization. Protein surface characterization was performed to evaluate the underlying chemical properties of the nanoparticle-binding regions on the ubiquitin surface. Electrostatic potential (EP) surface of ubiquitin was generated using the Adaptive Poisson−Boltzmann Solver (APBS) program. These maps have been used previously to provide insights into the charge interactions of proteins.48−51 For hydrophobic characterization, a spatial aggregation propensity (SAP) map was calculated, which provides information about the average hydrophobicity of each protein atom based on the hydrophobicity of the surrounding atoms and their surface exposure.52 This approach has been used to calculate hydrophobic patches on protein’s surfaces which can be used to predict aggregation hotspots and design proteins with enhanced stability.52,53

groups (Figure S1, Supporting Information). Further, particle size distribution was confirmed by performing transmission electron microscopy (Figure S2, Supporting Information). Reverse Phase Chromatography. The reaction mixture was loaded onto the C18 column in buffer A (DI water with 0.1% (v/v) TFA) followed by a step gradient to 40% buffer B (90% (v/v) acetonitrile in DI water with 0.1% (v/v) TFA) in order to elute the unreacted ligands. Following this, the final product was eluted with a step to 100% buffer B and collected. Isothermal Titration Calorimetry. A MicroCal VP-ITC system controlled by the VPViewer 2000 software was used to perform the titration experiments across a range of temperatures (5−35 °C). Each titration experiment consisted of 30−45 successive injections of a constant volume (10 μL) of ubiquitin solution (500−600 μM) into the sample cell (1.4 mL) which contained the NP solution (1.0−2.0 μM) in the same buffer (10 mM sodium acetate, pH 5.0, 0.02% (w/w) sodium azide). Blank injections of protein into the buffer were used to assess the heat of mixing and dilution and were subtracted from the heats of adsorption (Q). Q was obtained by integrating the peak after each injection using the ORIGIN software provided by the manufacturer and was then normalized by the amount of protein being injected. The resulting data were fit to an N independent sites binding model where the heat of adsorption is related to the calorimetric enthalpy of binding (ΔH), the stoichiometry of protein binding to the functionalized NP (N), and the dissociation constant (KD) of the protein−NP complex as shown in eqs 1 and 244

Q=

N[NP]0 ΔHV0 ⎡ ⎢A − ⎢⎣ 2

A=1+

A2 −

[P]0 K [P] + D 0 N[NP]0 N[NP]0

4[P]0 ⎤ ⎥ N[NP]0 ⎥⎦

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RESULTS AND DISCUSSION Multi Modal Ligand Functionalized Nanoparticles. As described in the Experimental Section, MM SAMs were synthesized to produce a pseudo-solid-phase system which qualitatively mimicked MM chromatographic resin materials (Figure 1A). This was done to facilitate the investigation of protein interactions with MM surfaces which have been shown to have unique selectivities for complex protein separations. The MM cation exchange ligands employed in this study were selected to be representative of two industrially important MM ligands: Capto MMC from GE Healthcare and Nuvia cPrime from Bio-Rad Laboratories (Figure S3, Supporting Information). While the “Nuvia ligand” is essentially identical to the ligand employed in the commercial resin material, the “Capto ligand” is missing a thioether moiety at the point of attachment. As described in the Experimental Section, the ligands used in this study were immobilized using an activated ester to attach the ligand headgroup to the linker, followed by attachment of the linker to the gold nanoparticles (NPs) via the thiol moiety through a place-exchange process.40−43 A schematic of the MM ligand functionalized NP surface is shown in Figure 1B,C. As can been in the figure, the major difference between both surfaces is the exposed aromatic group for the “Capto ligand” system as compared to the geometrically inaccessible aromatic group for the “Nuvia ligand” system. Further, the more linear structure of the “Nuvia ligand” along with its ability to efficiently π-stack could result in a more aligned ligand surface with the charged moiety facing out into solution. As described in the Supporting Information, proton NMR was employed to determine the ligand densities of the two MM-NP systems. The “Nuvia ligand” based NP had a ligand density of 2.5 ligands/ nm2 as compared to the “Capto ligand” system, which had a surface ligand density of 2.3 ligands/nm2 (see Supporting Information). The higher ligand density of the Nuvia system is not surprising since less steric hindrance would be expected for the formation of this MM SAM as compared to the more bulkier Capto ligand headgroup. Spacer arms are commonly employed in chromatographic systems to make the ligand more accessible and to minimize nonspecific adsorption to the base polymer matrix. Accordingly, a PEG linker with six monomer units was employed in our systems to prevent nonspecific binding of proteins to the underlying hydrophobic decane surface.39,40,54 The surface ligand densities (∼3 ligands/nm2) associated with the functionalized NP sytems are significantly higher than those commonly

(1)

(2)

where [P]0 and [NP]0 are the total protein and NP concentrations, respectively, in the ITC cell of volume V0. Protein Expression and Purification. Uniformly 15N-labeled wild-type recombinant ubiquitin (pI = 6.79, 8.5 kDa) was expressed using the BL21(DE3) strain of E. coli and purified as described by Chung et al.12 The final protein concentration was determined by spectrophotometric analysis at 280 nm with a molar extinction coefficient of 1490 M−1 cm−1.45 NMR Experiments. All NMR spectra were obtained at 35 °C using a Bruker 600-MHz spectrometer equipped with a 1H/13C/15N cryoprobe and z-axis gradients. Data were acquired and processed using Bruker TopSpin 2.1 software and the software package Sparky.46 Assignments of amide resonances were confirmed by matching published chemical shift values (BMRB accession number 6457) with backbone correlation patterns detected in spectra of unbound protein. 1H−15N HSQC spectra were acquired in a Shigemi 5 mm symmetrical NMR microtube assembly containing isotopically enriched 15N ubiquitin and functionalized NPs in NMR buffer (10 mM sodium acetate, pH 5.0, 0.02% (w/w) sodium azide, 150 μM TMSP as an internal standard, and 5% (v/v) D 2 O). The concentrations (μM) of NP and protein used in these experiments for the “Capto ligand” were 50, 40; 44, 80; 34, 160; 12, 320; and 0, 409.3, and those for the “Nuvia ligand” were 50, 77; 48, 115; 41, 154; 27, 231; 15, 308; and 0, 409.3, respectively. Experiments were performed at each of these conditions, and the resulting protein spectra were acquired. The maximum change in weighted average chemical shift (i.e., combined chemical shift) ΔδNH = ((ΔδH)2 + (0.2 × ΔδN)2)0.5 was calculated for each amide group that was observed to undergo ligand-induced changes in chemical shift.44,45 Finally curve fitting and calculations were performed with Origin 8 (OriginLab), and protein visualization was carried out with PyMol.47 Dynamic Light Scattering. Dynamic light scattering (DLS) was performed using a DynaPro Titan light-scattering apparatus (Wyatt, Santa Barbara, CA). The functionalized NPs were dissolved in a 10 mM sodium acetate buffer (pH 5.0 with 0.02% (w/w) sodium azide) at a concentration of 2.0 μM and then transferred into disposable lightscattering cuvettes in the absence or in the presence of ubiquitin. DLS D


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encountered in chromatographic resin systems (∼1 ligand/ nm2).55 DLS measurements of the NPs were carried out in the absence and presence of ubiquitin, wherein the ubiquitin concentration was maintained at a molar ratio of 50:1 to the NPs in order to ensure complete saturation of the NP surface. The average hydrodynamic radii for the NPs in the absence and presence of ubiquitin were found to be 1.4 ± 0.2 and 3.4 ± 0.3 nm, respectively. In addition, no larger species were detected. Further, a polydispersity of 14% was observed for the NPs in the presence of ubiquitin. Minimal if any polydispersity was observed for the pure NP solution. These results indicated that the NPs were stable and that no aggregation occurred for either NP system. Isothermal Titration Calorimetry. ITC has been frequently used to probe biomacromolecular interactions, directly providing the enthalpy of interaction, the stoichiometry of binding, and the dissociation constant of the binding process.39,56−59 ITC was employed in this study to investigate the energetics of binding between ubiquitin and the MM ligand functionalized NP surfaces for a range of temperatures (5−35 °C). As described in the Experimental Section, the protein ubiquitin was titrated into the sample cell containing the functionalized NPs, and the resulting heat changes were fit to an N independent sites binding model to quantify the corresponding thermodynamic parameters of protein−NP interactions. Figure 2A shows representative results of calorimetric traces in which ubiquitin was titrated into the sample cell containing the “Nuvia ligand” functionalized NPs at 5 °C. As can be seen in the figure, the heat of the interaction at this temperature was endothermic. The resultant fit to the N independent sites binding model (eqs 1 and 2) is presented in Figure 2B. The dissociation constants (KD), enthalpy changes (ΔH), and binding stoichiometries (N) were determined from the curvefitting analysis as described in Experimental Section and are summarized in Table 1. An experimental error of 14% was observed for values of ΔH. As can be seen in Table 1, the stoichiometry of protein binding to the NPs obtained with both systems over a range of temperatures was approximately 5 proteins per particle (the average N for the “Capto ligand” and “Nuvia ligand” systems were 5.1 ± 0.6 and 5.0 ± 0.2, respectively). This similarity of stoichiometry is not surprising since the dimensions of the two NP systems are expected to be quite similar and the maximum binding capacity is often a geometrically governed phenomenon. Of course, this would not be the case if the protein was not globular and if the protein had a different preferred binding orientation for the two systems. Since the N values were quite similar for these studies, the use of a constant value of 5 proteins per particle was employed for the calculation of residue specific dissociation constants in the NMR experiments described below. A rough geometrical calculation suggests that the accessible surface area (ASA) for the MM NP is ∼25 nm2 (considered as a sphere with a diameter of 2.8 nm as calculated from DLS experiments). The projected area of the ubiquitin molecule is ∼3 nm2 (considering ubiquitin as a sphere of diameter 2.0 nm as calculated from DLS experiments). This indicates that steric/geometrical limitations are limiting the number of bound proteins to roughly 60% of the accessible surface area. This is not surprising since ubiquitin is a small globular, relatively “hard” protein which is unlikely to undergo conformational changes which would be required to cover the

Figure 2. (A) Representative calorimetric traces of injection of ubiquitin into (a) buffer and (b) “Nuvia ligand” functionalized NP solution at 5 °C. (B) Dependence of the heat of ubiquitin−NP interaction on the [protein]/[particle] molar ratio. The curve represents the resulting fit using the N independent sites binding model (eqs 1 and 2).

Table 1. Dissociation Constants (KD), Enthalpy Changes (ΔH), and Binding Stoichiometries (N) at Different Temperatures for the Ubiquitin−MM Nanoparticle System “Capto ligand” T (°C) N (sites) ΔH (cal/mol) KD (μM)

“Nuvia ligand”

5 4.6 3562

15 5.1 2887

25 5.7 1251

5 4.9 2628

15 4.9 1895

25 5.2 −1160

35 5.0 −889

9

9

8

9

6

4

6

NP surface without steric limitations. In addition, lateral repulsions between bound proteins could also play a role in the reduced coverage. Further, it was also observed that ubiquitin interacts with both ligands with very similar affinities (the average KD for the “Capto ligand” and the “Nuvia ligand” was 9 ± 1 and 6 ± 2 μM, respectively). To gain further insight into the driving forces governing the ubiquitin−NP interaction, the Gibbs free energy changes (ΔG) and entropy changes (ΔS) were calculated using standard thermodynamic equations ΔG = −RT ln Keq and ΔG = ΔH − TΔS, and the resulting data from this analysis are presented in Figure 3. As can be seen in Figure 3, the titration experiments at different temperatures indicated minimal dependence of ΔG on the temperature for both ligand systems. In contrast, the ΔH values were functions of temperature for both ligands, with unfavorable enthalpy changes (ΔH > 0) observed for the “Capto ligand” for temperatures up to 25 °C (note: the value of ΔH decreased to zero at 35 °C, making data nondeterminable E


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desolvation enthalpy and favorable intrinsic enthalpy. Since the ΔH values were positive for majority of the data (Figure 3), it appears that desolvation was the major contributor to the enthalpy of binding. As shown in Figure 3 and Figure S4 (Supporting Information), the data suggest that the calorimetric enthalpy decreased with an increase in temperature for both ligand surfaces. Further, the heat capacity change of the binding interaction (ΔCp) obtained from the slope of Figure S4 was found to be negative for both ligand systems (−116 ± 28 cal mol−1 K−1 and −136 ± 43 cal mol−1 K−1 for “the Capto ligand” and “the Nuvia ligand”, respectively). The negative values of ΔCp in these systems are consistent with the process of dehydration of interfaces that occurs during binding.61 It is also important to note that only the release of loosely associated waters will have a direct effect on ΔCp. Thus, the relatively modest negative values of ΔCp reported in the paper (Figure S4, Supporting Information) are consistent with relatively small hydrophobic areas of interaction on the protein surface (which will be discussed below). Further, this does not account for the large entropic contribution that will come from the release of strongly associated waters from the charged surfaces which will not be reflected in the ΔCp values. Previous work has shown that protein binding in hydrophobic interaction chromatographic (HIC) systems is associated with entropically driven processes and that the magnitude of the entropic contribution increased with increased hydrophobicity of the ligand surface.58,59 Thus, the results presented in our paper suggest that from a thermodynamic perspective binding in the MM system is similar to HIC where entropic effects dominate. Nuclear Magnetic Resonance Spectroscopy. To facilitate further residue level investigation of protein−MM surface interactions, NMR spectroscopy was employed using functionalized NPs that simulate a chromatographic resin surface while also being amenable to solution NMR. To study the interaction of ubiquitin with the MM ligand surface, stoichiometric amounts of 15N-labeled ubiquitin was titrated against a fixed amount of MM ligand functionalized NPs in solution. The titrations were monitored through the 1H−15N HSQC spectra of ubiquitin as described in the Experimental Section. Briefly, in the NMR titration experiments, residues on the 15N-labeled amide backbone of the protein that came into close proximity with the MM surface experienced a change in the local electronic environment, resulting in a chemical shift perturbation (CSP). The amide groups thus served as reporters of the microenvironment for each residue on the protein. In the resulting 2D 1H−15N HSQC spectra, a single resonance peak was observed for both the unbound and the bound state of the protein for each amide group. As a result, for every point during the titration, the observed peak position in the spectra were observed to be at a population averaged chemical shift of bound and unbound protein states as given in eq 363,64

Figure 3. Comparison of the enthalpic (ΔH) and entropic (−TΔS) contributions to the Gibbs free energy (ΔG) of ubiquitin−NP interaction at different temperatures. Data are presented for both ligand systems.

at that temperature). In contrast, while the “Nuvia ligand” system exhibited an unfavorable enthalpy change at temperatures of 5 and 15 °C, a favorable value (ΔH < 0) was obtained at temperatures of 25 and 35 °C. An important result from these studies was that the binding of ubiquitin to both ligand surfaces at all temperatures was dominated by a large favorable entropy change (ΔS > 0), indicating that the binding of ubiquitin was entropically driven in these systems. Although protein binding to a MM functionalized NP can be quite complicated, it primarily consists of two simultaneous processes: (a) the intrinsic binding of the protein to the surface due to a combination of electrostatic, hydrogen bonding, π−π, and π−cation interactions and (b) desolvation of the NP and the protein and solvation of the newly formed protein−NP complex.60 Thus, the process of protein binding can be summarized by the following equilibrium between the protein and the NP as well as their solvated state. [protein]·m[H 2O] + [NP]·n[H 2O] + (q)[H 2O] ⇌ [protein·NP]·(m + n − x)[H 2O] + (x + q)[H 2O]

where m and n connote the number of water molecules solvating the protein and NP, respectively, and x is the number of water molecules released upon binding. Entropic changes (TΔS) can be broken down into two terms. First, there can be an unfavorable entropic contribution due to conformational restriction of flexible amino acid residues on the protein surface and the ligand head groups upon binding, i.e., a configurational entropic contribution. Second, there can be a favorable entropic contribution due to the water release that occurs during binding. Thus, for both ligand systems examined in this study, the large positive entropic gain observed across a range of temperatures indicates that the release of waters of hydration from the binding interfaces was the primary process governing these interactions.39,58,61,62 The enthalpy changes (ΔH) can be thought of as consisting of the intrinsic enthalpy of noncovalent bond formation which is exothermic (ΔHprotein−NP < 0) along with the dissociation of well-ordered waters of hydration which is endothermic (ΔHdesolvation > 0). Thus, the enthalpy changes observed in these experiments are overall outcomes of unfavorable

δobs = x uδu + xbδ b

(3)

where xu and xb are the mole fractions of unbound and bound protein, respectively, and δu and δb are the chemical shifts of the unbound and bound states. Since titrations were carried out by the addition of ubiquitin to a fixed mass of NPs, during the initial titrations, the NPs were in excess resulting in the majority of the proteins in the bound state. As the titrations continued, the average population F


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Figure 4. Ligand-induced combined chemical shift changes for (A) “Capto ligand” and (B) “Nuvia ligand” across all ubiquitin residues.

of proteins moved toward the unbound state. A steady decrease for the vast majority of the 1H−15N signals in the HSQC spectra was observed as the system moved toward higher NP to protein ratios, eventually leading to complete disappearance of these signals at ratios exceeding 2:1. Extensive broadening or loss of NMR signals can likely be attributed to the following phenomena: an intermediate chemical exchange due to interconversion of the free and bound states of ubiquitin (i.e., exchange broadening) and/or the large molecular size of the resulting complex in solution.48,49 For intermediate chemical exchanges, the line width is dependent on the exchange kinetics, and a slower kinetic regime results into broader spectra.64 Since exchange kinetics are a strong function of temperature, a corresponding increase in temperature should result in decreased peak width (i.e., sharper spectra).65 In addition, as the molecular size of the complex increases, the line width (i.e., short transverse relaxation time, T2) of the signal increases.57 Increasing the temperature for such a system causes the NP−protein complex to tumble faster in solution, leading to a reduction in line width and, hence, sharper spectra. Since increasing the temperature would enhance the signal-to-noise ratio for both scenarios, all NMR titrations were carried out at temperature of 35 °C. As a result of this higher temperature, an overall decrease in peak widths (i.e., sharper spectra) was observed with increasing NP concentration, which is consistent with exchange broadening being a contributor to peak disappearances. However, even at this temperature, the larger molecular size of the protein−NP complex still gave rise to some broadening at higher NP to protein ratios. It is important to observe the distribution of the CSPs across the entire protein surface as part of an initial analysis. A plot of the maximum observed CSP (Δδobs) obtained during titration for each residue is shown in Figure 4. Although CSP data facilitate understanding of protein−ligand interactions, the interpretation is not trivial. CSPs can occur for residues that are not directly involved in binding but are in close proximity to the binding site. Also, a residue directly interacting with the ligand may not necessarily undergo a significant CSP depending on microenvironment effects. To overcome these ambiguities and to gain quantitative information from these spectra, the changes in chemical shift during the titration experiments were examined in further detail. Chemical shift changes were monitored through the 2D 1 H−15N HSQC spectra. Figure 5 shows representative HSQC spectra for some of the residues that experienced ligand-

Figure 5. Representative 1H−15N HSQC spectra from the “Capto ligand” titration experiments at 0 μM (magenta), 12 μM (red), 34 μM (green), 44 μM (purple), and 50 μM (blue) NP concentrations: (A) 69 leucine, (B) 48 lysine, and (C) 68 histidine.

induced changes in chemical shift and trended toward saturation. The majority of the observed changes in chemical shift suggested a simple two-state binding process. Figure 5A and 5B for residues 69L and 48K, respectively, are representative HSQC spectra that show such a two-state behavior. Only one residue, 68H, clearly exhibited a complex G


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Figure 6. Multimodal ligand surface binding residues determined by NMR with color-coded dissociation constants for (A) “Capto ligand” surface and (B) “Nuvia ligand” surface. (C) SAP map and (D) EP map for ubiquitin.

ligands). The assumptions of the model are (i) all potential N binding sites on the functionalized NP are equivalent and (ii) the same value of stoichiometry (N) (i.e., the maximum number of available sites on the NP) can be employed when fitting the model for each residue. The value of N employed in this analysis was determined to be 5 proteins per NP from independent ITC experiments carried out on the same protein NP system discussed in the previous section. The dissociation constants (KD) and the maximum attainable change in chemical shift (Δδmax) for each interacting residue on the protein surface were determined by fitting the model (eqs 4−9) using a stoichiometry of N = 5. As shown by the representative results illustrated for several residues (Figure S5, Supporting Information), good agreement is seen among fits of the model and the experimental data. A maximum fitting error of 20% was obtained with a 95% confidence interval for the fits of the individual residue dissociation constants (KD). While residues with significant interactions with the MMNPs exhibited well behaved trends in the chemical shift change as shown in the representative shifts in Figure S5, residues with maximum chemical shift values of 0.09 ppm and below had significant noise in the data indicating nonspecific interactions. Thus, these residues with low chemical shifts were not included in our analysis to minimize the contribution of nonspecific interactions. Figure 6A,B shows the location of residues on the protein surface that had measurable KD values for each ligand system.

multistate binding behavior resulting in a nonlinear trajectory in frequency space, as shown in Figure 5C. For residues that showed significant CSPs, the observed chemical shift changes (Δδobs) were fit to an N independent sites binding model (eqs 4−9) to obtain residue specific dissociation constants (KD) and maximum chemical shifts (Δδmax). [P] + [S] ⇌ [P·S] KD =

([S]0 − [P·S])([P]0 − [P·S]) [P·S]

[S]0 = N[NP]0

(4)

(5) (6)

[P·S] = (1/2)[(KD + N[NP]0 + [P]0 ) −

(KD + N[NP]0 + [P]0 )2 − 4N[NP]0 [P]0 ) ]

Δδobs = Δδmax([P·S]/[P]0 )

(7) (8)

Δδobs = (Δδmax /2[P]0 )[(KD + N[NP]0 + [P]0 ) −

(KD + N[NP]0 + [P]0 )2 − 4N[NP]0 [P]0 ) ]

(9)

A binding site (S) in this context corresponds to the interface on the functionalized NP where a single protein (P) will bind (note: such an interface will likely include multiple MM H


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The KD values are presented in a color scale ranging from red (10 μM) to blue (500 μM) with residues that did not exhibit significant CSPs indicated in black. As can be seen in the figure, the majority of the residues involved in binding were distributed across the front face of ubiquitin for both ligand systems. For the “Capto ligand” NP system (Figure 6A), it can be observed that the residues on the protein with the strongest interactions were localized around the group of aliphatic residues with the possible exception of a single interacting residue on the back face of the protein (32D). The residues involved in binding on the front face were distributed around the strongest binding residue 8L (16 μM). Thus, the “Capto ligand” seems to selectively bind to a specific cluster of aliphatic residues on the front face of ubiquitin. On the other hand, for the “Nuvia ligand” NP system (Figure 6B), a more diffuse type of binding behavior was observed which included a contiguous binding region on the front face, composed of an aliphatic residue region (69L, 70 V, and 71L) and polar residues (9T, 10G, 47G, 72R, 74R, 68H, 6K, and 48K). Moreover, the “Nuvia ligand” NP system also exhibited moderate interactions with the back face of ubiquitin (25N). These results suggest that the two MM NP systems have very different modes of interactions with ubiquitin. These findings are quite interesting since the two ligands have very similar chemistries with subtle differences in their geometrical presentation of the ligands. As was indicated above in the discussion related to Figure 1, the major difference between the two MM NP surfaces is the exposed aromatic group for the “Capto ligand” system as compared to the geometrically less accessible aromatic group for the “Nuvia ligand” system. In order to examine the relationship between protein surface properties and this NMR data with the two ligand systems, we carried out two analyses. The electrostatic potential (EP) across the protein surface was first determined to examine protein surface charge effects (Figure 6D). As can be seen in the figure, this EP map indicates the presence of a strong positively charged contiguous surface across the front face of the protein (encircled in yellow). The region encircled in yellow in Figure 6D includes several positively charged residues on the front face of the protein that were also identified in the NMR experiment with the “Nuvia ligand” (72R, 74R, 68H, 6K, and 48K). Interestingly, this EP surface corresponds better to the NMR results obtained with the “Nuvia ligand” NP system (Figure 6B) as compared to the “Capto ligand” NP system (Figure 6A). As discussed previously in the Results and Discussion section, the aromatic moiety on the immobilized Nuvia ligand NP is likely more sterically hindered which will potentially increase the relative role of electrostatic interactions with this surface. The spatial aggregation propensity (SAP) was then determined to examine the hydrophobic surface properties of ubiquitin and a SAP map is presented in Figure 6C.52 As can be seen in the figure, using this analysis a cluster of aliphatic residues (8L, 71L, and 69L) on the front face of ubiquitin produced a strongly hydrophobic region (dark red) on the protein surface (encircled in black). Interestingly, this cluster of aliphatic residues (8L, 71L, and 69L) was also identified in the NMR experiments with the “Capto ligand” NP system as being a high binding region (Figure 6A), thus supporting the hypothesis that the exposed aromatic group for the “Capto ligand” has a strong effect on the selective binding of the ligand groups to the hydrophobic residues on the protein surface. It was also of interest to examine the overlap or proximity of the defined EP and SAP regions. The front face of the protein

possesses both positive EP (Figure 6D) and hydrophobic SAP (Figure 6C) regions. On the other hand, the back face of the protein possesses both negative EP and hydrophilic regions. Moreover, these regions on the front and back faces of the protein have significant overlap. Thus, it is not surprising that ubiquitin has a preferred binding region on the front face as compared to the back face. Importantly, this is supported by the NMR data which demonstrate a preferred binding face with both MM ligand systems. It is also worth noting that in multimodal chromatography where both electrostatic and hydrophobic interactions can play an important role different MM ligand systems will vary in the relative importance of these two contributions as is clearly evidenced by this NMR data. It is of importance to note that the residue specific binding constants obtained for the “Capto ligand” NP system (μM affinity) are several orders of magnitude stronger than those previously obtained with ligands in free solution (mM affinity).12 This supports the hypothesis that the cluster of interacting residues identified on the protein surface act cooperatively to simultaneously occupy a “binding site” on the multimodal surface which consists of several multimodal ligands. Further, it is interesting to note that the binding affinities for overall protein−surface interaction obtained from the ITC experiments (Table 1) were in qualitative agreement with the residue level dissociation constants measured from the NMR chemical shift experiments that showed both MM-NP surfaces interacting with the high interaction protein residues in the low μM range (Figure 6). The results from ITC suggested that from a thermodynamic perspective, binding in the MM system is similar to HIC where entropic effects dominate. The importance of hydrophobic interactions in these MM NP systems was also highlighted by the NMR studies through the identification of the specific residues composing the interaction surfaces. Recent work has indicated that hydrophobically driven binding of solutes in the small length scale regime (less than 1 nm2) will be entropic in nature.66−68 While the NMR determined surface patches on ubiquitin involved in binding consisted of both hydrophobic and polar residues, the hydrophobic patches were all less than 1 nm2 (Figure 6A,B). Thus, the entropic nature of binding is also consistent with this vision of the thermodynamics of hydrophobic interactions.

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CONCLUSION In this paper, we have developed and demonstrated the use of a novel pseudo-solid-state system to investigate protein interactions with two different MM chromatographic ligand surfaces. The nature of binding of ubiquitin to MM ligand functionalized NPs was examined using ITC and NMR. While the SAM-based MM−NP system may not be an exact representation of MM chromatography due to a more heterogeneous distribution of ligands on the resin, we believe that the use of MM−NPs can provide important qualitative insights into the effects of MM ligand chemistry and presentation on protein adsorption behavior. ITC was performed on the ubiquitin−NP system over a range of temperatures to obtain insights into the thermodynamic driving forces governing the protein−MM surface interactions and to measure the stoichiometry (N) of the binding process. ITC revealed that binding to the MM ligand surface was entropically driven at all temperatures and that this involved the release of surface bound waters upon complexation. This behavior was true for both MM ligand systems. The entropic nature of I


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chromatographic ligand structures of Capto MMC from GE Healthcare and Nuvia cPrime from Bio-Rad Laboratories (Figure S3), temperature dependence of ΔH of interaction for ubiquitin with the NPs and linear fit of the data to obtain ΔCp (Figure S4), representative ligand-induced chemical shift change data and resulting fits to the N independent site model (Figure S5). This material is available free of charge via the Internet at http://pubs.acs.org.

binding observed from ITC combined with previous work on HIC systems in the literature suggests that, from a thermodynamic perspective, MM systems may behave more like HIC than ion exchange.69−71 It is interesting to note that the hydrophobic patches on the surface of ubiquitin involved in binding fall in the regime of “small length scale” hydrophobicity (less than 1 nm2), which has been shown to be entropic in contrast to larger length scale hydrophobicities.66−68 This is in concert with the entropic driving force obtained from the ITC experiments. Further, to investigate the binding processes at the residue level, NMR experiments with 15N-labeled ubiquitin were performed, which revealed the presence of a preferred binding face on ubiquitin for interaction to both MM ligand surfaces. The dissociation constants obtained from ITC were in qualitative agreement with those obtained for individual amino acid residues from the NMR experiments. The residue level dissociation constants suggested that binding of ubiquitin to the “Capto ligand” was more specific and localized to an aliphatic residue region on the protein surface whereas binding to the “Nuvia ligand” was more diffuse, involving several high binding residues across the interface. Further, the cluster of aliphatic residues (8L, 71L, and 69L) identified by NMR measurements with the “Capto ligand” surface (Figure 6A) coincided well with the same group of residues identified by the SAP map (Figure 6C).52 Comparison of the results from the NMR experiments (Figure 6A,B) and the surface characterization maps (Figure 6C,D) suggests that although both ligand systems have an overlap of hydrophobic and electrostatic regions across the binding face of the protein, in comparison to the “Capto ligand”, the interaction of the “Nuvia ligand” system has a more substantial contribution from electrostatic interactions. The back face of the protein showed a significant overlap of both negative EP and hydrophilic regions which explain the absence of significant binding regions on this face. The present work highlights the importance of co-operativity associated with protein−surface interactions since the cluster of interacting residues on the protein surface act coherently to simultaneously occupy a binding site on the NP surface and as a result give rise to MM binding affinities several orders of magnitude greater than those obtained for interactions with free solution ligands.12 Although the focus of this work has been on human ubiquitin, this approach should be readily applicable to other protein−NP systems provided that the relative sizes of the proteins and NPs are comparable. It would also be of interest to examine the behavior of larger proteins which may have a less focused interaction with the multimodal surface. Ligand density has been shown to have a significant effect on protein separation selectivity.71,72 The protein−NP system presented in this work provides a powerful platform for controlling and studying ligand density effects at the molecular level. Accordingly, future work will employ the MM−NPs to examine how ligand density affects co-operativity and avidity in these MM systems. Future work will also examine alternative chromatographic ligand systems ranging from pseudoaffinities to high affinities such as immobilized peptides.73

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected] (S.M.C.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by National Science Foundation Grant CBET 1134341. We thank Lars Sejergaard for assistance with the NMR data analysis and Blanca Barquera for assistance with the expression of the isotopically enriched ubiquitin.

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REFERENCES

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ASSOCIATED CONTENT

S Supporting Information *

Calculations for nanoparticle surface ligand density through quantitative proton NMR; 1H NMR spectrum of functionalized nanoparticles (Figure S1), TEM image of functionalized nanoparticles with particle size distribution (Figure S2), J


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