# Identification of Ni2++-binding peptides in sunflower meal protein hydrolysate for deeper understanding of peptide-metal interactions

Jairo Andrés Camaño Echavarría, Christelle Mathé, Jean-Michel Girardet, Cédric Paris, Chibuike Udenigwe, Katalin Selmeczi, Laetitia Canabady-Rochelle

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## ► To cite this version:

Jairo Andrés Camaño Echavarría, Christelle Mathé, Jean-Michel Girardet, Cédric Paris, Chibuike Udenigwe, et al.. Identification of Ni2++-binding peptides in sunflower meal protein hydrolysate for deeper understanding of peptide-metal interactions. Journal of Inorganic Biochemistry, 2025, 269, pp.112877. 10.1016/j.jinorgbio.2025.112877. hal-05151463

HAL Id: hal-05151463

https://hal.univ-lorraine.fr/hal-05151463v1

Submitted on 8 Jul 2025

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Distributed under a Creative Commons Attribution 4.0 International License
Sunflower meal by-product has widely been used to produce bioactive peptides. Naturally encrypted within protein sequence under an inactive form, these bioactive peptides are released upon proteolysis. Recently, sunflower meal-derived peptides have gained significant attention due to their wide-ranging health benefits such as immunomodulators [3], angiotensin-converting inhibitors [4], antioxidants, anti-inflammatory [5] and metal-binding compounds [6].

Metal-binding peptides can be powerful ligands to complex several transition metal ions such as $Ni^{2+}$, $Cu^{2+}$, $Zn^{2+}$ and $Fe^{2+}$/$^{3+}$. These peptides have been proposed as a powerful substitute for metals-related diseases [7]. For instance, through the complexation of $Fe^{2+}$ and $Cu^{2+}$, they may effectively delay oxidative damage linked to chronic diseases [8]. Besides, metal-binding peptide can promote better stability, absorption, and bioavailability of minerals and are endowed with important biological functions [9,10]. Metal-binding peptides can be released from various protein sources such as milk, eggs, and fish [7]. Nonetheless, from the standpoint of sustainable development and the utilization of biological resources, numerous studies have explored the potential of agro-industrial by-products, given their substantial protein content and their added value once hydrolyzed [5]. Although several metal-binding peptides have been reported from agro-industrial by-product-derived proteins (walnut flake [11], chia seed [12]), the evaluation of sunflower meal by-product remains scarce [6,13,14].

Nickel is important for many microorganisms and is a component of several metalloenzymes [15]. Although it is not an essential metal for human life, nickel can serve as a probe to better understand the peptide-metal interactions along with other metal ions (e.g., $Cu^{2+}$). Indeed, many similarities have already been described between $Ni^{2+}$ and $Cu^{2+}$ ion complexing processes [16]. Besides, immobilized metal ion affinity chromatography (IMAC) is a purification technique based on the affinity of some peculiar amino acids (AA) residues present in peptides for some transition metal ions such as $Ni^{2+}$ immobilized onto the chromatographic solid phase. Thus, $Ni^{2+}$-IMAC can be used both to separate and purify metal-binding peptides containing potential metal-binding AA, especially histidine residue, known to be an excellent anchoring site for $Ni^{2+}$ and $Cu^{2+}$ ions [17], and to study the $Ni^{2+}$-binding ability of His-containing peptides [18]. However, only few information is available on the degree of affinity between the His-containing peptides and the immobilized $Ni^{2+}$ ions. Recently, the switchSENSE® technology has been developed and successfully used to study in real-time peptide interactions with immobilized $Ni^{2+}$ ions strongly bound to the tetradentate nitrilotriacetic acid (NTA), while providing complementary and deeper information about the peptide-Ni$^{2+}$ complex [19]. Despite all the advantages of IMAC and switchSENSE® technology to study the Ni$^{2+}$ binding ability of food-derived peptides, the conformational geometry and binding modes of the peptide-Ni$^{2+}$ complexes should be further characterized to better understand this specific interaction. Spectroscopic techniques such as UV-visible (UV-vis) and circular dichroism (CD) provide some information on the coordination characteristics of peptides with Ni$^{2+}$ ions, revealing significant data about the molecular basis of the peptide-metal interaction such as binding sites and geometry of the metal ion environment in the formed complexes [15].

To the authors' knowledge, there is no report on Ni$^{2+}$-binding properties of His-containing peptides from sunflower meal by-product and the influence of the amino acid sequence on such biofunctionality. Here, single and sequential enzymatic hydrolysates were produced from sunflower meal protein isolate (SMPI) hydrolysed using either Protamex® (Prot) or Protamex followed by Flavourzyme® (Prot+Flav), respectively in order to study their Ni$^{2+}$-binding properties using IMAC, switchSENSE®, UV-visible and CD techniques. Additionally, ten pure peptide sequences theoretically present in sunflower proteins were investigated to better understand the formation of peptide-Ni$^{2+}$ complex. This paper focuses on how sunflower meal by-product may be used in the production of food-derived metal-binding peptides. Such knowledge (i) will be an important input to understand the interaction between the peptide sequences present in SMPI hydrolysates and Ni$^{2+}$

ions, (ii) and will explain the bioinorganic chemistry of the metal-binding mechanism of the formed sunflower peptide-Ni$^{2+}$ complexes.

## 2. Materials & methods

### 2.1. Chemicals

Sunflower meal (SM) was provided by SAS IMPROVE (Dury, France). Protamex® (Prot, protease from Bacillus sp., ≥ 1.5 U/g), Flavourzyme® (Flav, protease from Aspergillus oryzae, ≥ 500 U/g), hydrochloric acid (37 %, HCl), ammonium bicarbonate, ethylenediaminetetraacetic acid ($Na_2EDTA-2H_2O$), nickel(II) sulfate ($NiSO_4·xH_2O$) and imidazole were purchased from Sigma-Aldrich (St Louis, MO, USA). All the other reagents were analytical grade and purchased from VWR International (Atlanta, USA): sodium phosphate monobasic $NaH_2PO_4$, sodium phosphate dibasic $Na_2HPO_4·12H_2O$, sodium chloride, sodium hydroxide, HEPES (99 %, $C_8H_{18}N_2O_8S$).

### 2.2. Synthetic peptides investigated

Ten small-sized peptides ($≤ 10$ AA residues), naturally encrypted in the major sunflower protein (11S globulin seed storage protein G3, P19084, 11S3_HELAN) were selected; the sequence of this former protein was found in UniProt protein data base (https://www.uniprot.org/uniprotkb/P19084/entry) (Fig. S1). The peptides sequences were chosen according to their particular structures, which could confer them different affinity for Ni$^{2+}$ ions based on the presence of His, Trp, Cys, Glu, Asp and Pro residues (according to the Hard and Soft Acid Bases (HSAB) theory and Porath's triad, vide infra) and their position within the sequence. The peptides were classified into 3 groups: group (I) His or Trp-containing peptides without proline (Pep1-4), and it is divided into two subgroups: IA is composed by His-containing peptides (1. WLHNDGN, 2. WLHNDGNE, 3. FATCIARH), while the second one IB is composed by a Trp-containing peptide (4. WVSFKTN); group (II) His-containing peptides with proline (5. AIPGTAH, 6. KFPILEH, 7. KFPILEHL and 8. KFPILEHLR); and group (III) peptides without His or Trp (9. GNPO, 10.LLAF). The 6His peptide (11. HHHHHH) was also investigated as positive control. The peptide sequences selected were synthetized by Genecust (Dudelange, Luxemburg) and characterized by their molecular weight (MW), isoelectric point (pI, calculated from PepCalc free software, https://pepcalc.com) and their percentage of global charge at pH 7.4 using ACD/Percepta Software version 14.0 (ACD/Labs, https://www.acdlabs.com/). The chemical structures of the 11 synthetic peptides are presented in Fig. 1, while Table 1 shows their physicochemical properties.

### 2.3. Metal-binding peptides investigated in hydrolysate

#### 2.3.1. Hydrolysate production

#### 2.3.1.1. Sunflower protein isolate.
Sunflower meal protein isolate (SMPI) was obtained using the method reported by Kalpana et al. [20] based on NaCl extraction to solubilize the majority of globulins proteins. Sunflower meal (30 g) was dissolved in a 10 % (w/v) meal-to-ultrapure water ratio, added with 10 % (w/v) of NaCl. Then, this former solution was stirred on a magnetic stirrer plate (60 min, 40 °C, pH adjusted at 6). After solubilization, the sample was cloth filtrated in order to remove solid fibrous residues. The resulting solution was centrifuged (Centrifuge Universal 320, Hettich, Tuttlingen, Germany) at 3300g, for 10 min, at 20 °C. Afterward, the resulting supernatant composed of solubilized proteins was recovered and filtrated by vacuum filtration to remove the residues remaining from solubilization. The filtrated supernatant was precipitated within 10 min by adjusting the pH at 4.5, corresponding to the isoelectric point of sunflower proteins [13]. This suspension was let at rest for 30 min at room temperature (25 °C), and then centrifuged
<figure><img src="image_3.png" /><figcaption>**Fig. 1.** Chemical structures of pure peptides studied by IMAC, SwitchSENSE®, UV-visible and CD spectroscopic techniques. (A, B, C) His-containing peptides without proline (group I), (E, F, G, H) His-containing peptides with proline (group II), (D, I, J) No His-containing peptides, and (K) positive Ni²⁺-binding peptide (6His peptide). (L) schematic representation of ATCUN peptide-metal (Mn⁺) complex. Imidazole, indole, thiol and carboxylate groups could play a role in the Ni²⁺-binding properties (circle).</figcaption></figure>
<caption>**Table 1**</caption>

Pure peptides selected from 11S globulin protein sequence with their theoretical physico-chemical properties. (+) positive charge, (−) negative charge.

<table><thead><tr><th>Protein code</th><th>Protein name</th><th>Name</th><th>Peptide sequence</th><th>Length (AA)</th><th>Molecular mass (Da)</th><th>Isoelectric point (pI)</th><th>% of global charge at pH 7.4</th></tr></thead><tbody><tr><td rowspan="11">P19084</td><td rowspan="11">11S globulin seed storage protein G3</td><td>Pep1</td><td>WLHNDGN</td><td>7</td><td>854.35</td><td>4.87</td><td>0 % +, 86 % −</td></tr><tr><td>Pep2</td><td>WLHNDGNTE</td><td>9</td><td>1084.44</td><td>3.92</td><td>0 % +, 100 % −</td></tr><tr><td>Pep3</td><td>FATCIARH</td><td>8</td><td>918.08</td><td>8.62</td><td>75 % +, 2 % −</td></tr><tr><td>Pep4</td><td>WVSFKTN</td><td>7</td><td>880.99</td><td>10.01</td><td>43 % +, 0 % −</td></tr><tr><td>Pep5</td><td>AIPTGTAH</td><td>8</td><td>766.38</td><td>7.88</td><td>45 % +, 11 % −</td></tr><tr><td>Pep6</td><td>KFPILEH</td><td>7</td><td>882.48</td><td>7.60</td><td>26 % +, 20 % −</td></tr><tr><td>Pep7</td><td>KFPILEHL</td><td>8</td><td>995.57</td><td>7.60</td><td>18 % +, 33 % −</td></tr><tr><td>Pep8</td><td>KFPILEHLR</td><td>9</td><td>1151.67</td><td>9.90</td><td>64 % +, 0 % −</td></tr><tr><td>Pep9</td><td>GNPQ</td><td>4</td><td>414.17</td><td>3.70</td><td>0 % +, 34 % −</td></tr><tr><td>Pep10</td><td>LLAF</td><td>4</td><td>462.27</td><td>3.70</td><td>0 % +, 28 % −</td></tr><tr><td>Pep11</td><td>HHHHHH</td><td>6</td><td>840.85</td><td>7.97</td><td>70 % +, 3 % −</td></tr></tbody></table>

again (3300 g, 10 min, 20 °C). The pellet was finally dried using a convective oven (Thermo Scientific™, USA) at 55 °C overnight. SMPI was stored at −20 °C until further use.

### 2.3.1.2. Proteolysis.
Sunflower meal protein isolate (SMPI) was then hydrolysed as previously described by El Hajj et al. [21] with some modifications. First, 5 g of SMPI were dissolved in 250 mL of 50 mM ammonium bicarbonate buffer solution. Then, SMPI was hydrolysed either with Protamex® (Prot) or Protamex® followed by Flavourzyme® (Prot+Flav). The hydrolysis parameters were as follows: 2 % (w/v) SMPI concentration, 1 % (w/w) enzyme/substrate ratio, for 2 h, at pH 8 and 55 °C with Protamex® and for 2 h, pH 7 and 55 °C with Flavourzyme®, which correspond to their respective optimal conditions. Upon hydrolysis, the pH was maintained at the desired values by adding 1 M NaOH solution. After each hydrolysis step, each protease was heat inactivated (95 °C, 15 min). In the sequential treatment, Protamex® was inactivated just before Flavourzyme® treatment. Then, hydrolysates were centrifuged (10,000g, 15 min, 25 °C) in order to remove the enzymes, and the whole hydrolysates fractions were freeze-dried (Lab-conco, Kansas City, USA) and stored at −20 °C for further analysis.

These two hydrolysate powders were further dissolved (quantity sufficient for (qsf) 250 mL; 2 % (w/v) in ultrapure water) and submitted to ultrafiltration under magnetic stirring and 5 bars of nitrogen pressure [21]. The ultrafiltration unit (UF stirred cell 47 mm system XFUF04701; Millipore, Molsheim, France) was initially equipped with a 10 kDa molecular weight cut-off (MWCO) membrane, followed by a 1 kDa MWCO membrane (Millipore, Billerica, USA), conditioned according to their specification, before ultrafiltration. The ultrafiltrated fractions of hydrolysates (the so-called ≤ 1 kDa Prot and ≤ 1 kDa Prot+Flav) were freeze-dried and kept at −20 °C until further analysis. On total, four hydrolysates were investigated, whole hydrolysates: Prot and Prot+Flav, and ≤ 1 kDa fraction hydrolysates: ≤ 1 kDa Prot and ≤ 1 kDa Prot+Flav.

### 2.3.2. Immobilized metal ion affinity chromatography

The eleven synthetic peptides and the Ni²⁺-binding peptides present in whole hydrolysates and their ≤ 1 kDa fractions were investigated by Ni²⁺-IMAC using a HPLC system (LC-20AD, Shimadzu, Japan) at 1 mL/min flow rate, an oven temperature set at 25 °C and a diode array detector (SPD-M20A, Shimadzu, Japan). The Ni²⁺-binding peptides were separated on a 1 mL HisTrap™HP column (7 × 250 mm) prepacked with Ni Sepharose High Performance (Cytiva, Marlborough, MA, USA). Experiments were carried out according to Irankunda et al. [18] with some modifications. The method included five steps: nickel(II) regeneration, equilibration, injection followed by unbound compounds removal, elution and washing. First, 10 column volumes (CV) of ultrapure water were used to remove the ethanol remaining in the column (storage conditions). Then, the column was stripped using 50 mM EDTA solution, followed by nickel(II) regeneration step using a 100 mM nickel(II) sulfate (NiSO₄) solution. Then, 20 CV of equilibration buffer (PBS 1X pH 7.4: Na₂HPO₄ 6.7 mM; KH₂PO₄ 1.25 mM; NaCl 300 mM) were used to

remove the unbound nickel ions and equilibrate the column. Finally, 50 µL of either synthetic peptides (20 mM) or hydrolysates (20 mM eq. NH₂ according to OPA analysis, Table S1) were loaded onto the Ni²⁺-IMAC column (dead time of the column and the whole HPLC system: 1.2 min) [18]. The first fraction (F1) corresponding to the non-nickel-binding peptides was eluted with the equilibration buffer after 5 min (retention time (RT) comprised between 0 and 5 min), then the Ni²⁺-binding peptides fractions (F2) were eluted with an imidazole gradient (0–300 mM, prepared in PBS 1X pH 7.4) from 5 to 65 min (RT comprised between 5 and 65 min). Finally, 5 CV of a 600 mM imidazole solution was used to wash the columns (5 min) before the next run. Due to imidazole absorption at 214 nm, which is also the wavelength used for monitoring peptide bonds, the peptides were followed at 232 nm. For hydrolysates, the F1 and F2 fractions were collected using a fraction collector (FRC-10 A, Shimadzu, Japan) for further LC-MS/MS identification of peptide sequences. The retention time (RT) and the Imidazole concentration at which elution occurs (IMC) was determined for peptides and hydrolysates.

### 2.3.3. Metal-binding peptides identification by LC-MS/MS

#### 2.3.3.1. LC-MS/MS analysis.
To identify the peptide sequences present in the produced hydrolysates, the Ni²⁺-IMAC collected fractions (F1, F2) were evaluated on a Thermo Scientific UHPLC-HRMS system composed of a Vanquish™ liquid chromatography unit coupled to a photodiode array detector and an Orbitrap ID-X™ Tribrid™ high-resolution mass spectrometer operating in positive electrospray ionization mode (ESI+) as previously reported [22].

#### 2.3.3.2. Peptide sequence identification.
The Thermo XCalibur raw LC-MS/MS data files resulting from the collected fractions (F1, F2) from the hydrolysates in Ni²⁺-IMAC were processed in the Andromeda search engine of the commonly used proteomic software tools MaxQuant™ (version 2.4.13.0) [23]. The peptide sequences were searched against a restricted sunflower seed proteins data base (Helianthus annuus) containing 30 entries obtained from UniProtKB (https://www.uniprot.org/, accessed on 27 January 2024, Supplementary data, Table S2). The peptides identification was achieved by comparing the experimental fragment-ion mass value of the precursor ions with the theoretical fragment masses obtained from the protein sequences data base, obtained by the unspecified digestion rule with a mass tolerance of 10 ppm. The parameters of database searches were set up according to Boachie et al. [23] (Table S3). Note that before searching peptides sequences from the collected fraction of hydrolysates, this method was first validated using a synthetic peptides hydrolysate composed of 8 pure peptides at 1 mM each (Pep1–8, section 2.2). All these 8 peptides were clearly identified (Fig. S2). Finally, the AA composition of identified Ni²⁺-binding peptides present in SMPI hydrolysates (F2) were computed through ExPASy ProtParam server (https://web.expasy.org/protparam/).
## 2.4. Metal-peptides interaction

### 2.4.1. *switchSENSE®* technology

The study of the interaction between Ni²⁺ ions and peptides (either present in synthetic peptides or SMPI hydrolysates) was carried out with a biosensor analyzer DRX using an electrically switchable DNA chip designated as MPC-48-2-R1-S, incorporated NTA and Cy5 dye was used as a fluorescent probe (switchSENSE® technology; Dynamic Biosensors GmbH, Munich, Germany). This technology can be used both in static and in dynamic modes. The principles of these modes are reported by Muller-Landau [24]. Experiments were conducted according to the method initiated by El Hajj et al. [19] with some modifications. Briefly, the synthetic peptides (at 10 µM), the whole hydrolysates and their ≤ 1 kDa ultrafiltrates (at 200 µM eq. NH₂) were prepared using T40 buffer (i.e., 10 mM Tris-HCl, 40 mM NaCl, and 0.05 % v/v Tween 20 at pH 7.4). Then, the former sample solutions were injected into the microfluidic device at a flow rate of 30 µL/min for a 4 min association phase, followed by the injection of T40 buffer at 50 µL/min for a 4 min dissociation phase to release the analytes. After each investigated concentration (hydrolysates: 10, 100, 200 µM, synthetic peptides: 1, 5, 10 µM), Ni²⁺ ions were removed using a 100 mM EDTA solution, followed by a Ni²⁺ rebinding phase before launching another run. A buffer blank was also investigated to correct all kinetic experiments for non-specific interactions (static mode measurements).

The flow was paused between the association and the dissociation phases in static mode measurements in order to perform the dynamic mode measurement. This dynamic analysis enabled to evaluate the specificity of the peptide-metal ion interaction. The analysis of the different kinetic curves was performed using the SwitchANALYSIS® software from Dynamic Biosensors using non-linear fitting of single exponential functions. Each kinetic experiment was performed twice using different sample preparations to ensure reproducibility. For each experiment, changes in fluorescence (ΔF) and dynamic response (ΔDR) were measured and averaged over eight replicates obtained under identical conditions. Non-specific binding was corrected by subtracting the signals from the reference electrode (without NTA-Ni²⁺), and the resulting ΔF and ΔDR values, reflecting only specific binding, were reported as the final averages.

### 2.4.2. UV–visible spectroscopy

The determination of the UV-visible (UV-vis) spectrum of Ni²⁺ ions and peptide-Ni²⁺ complexes formed by synthetic peptides and SMPI hydrolysates (whole and ≤ 1 kDa fraction) was carried out at the PhotoNS Platform of the L2CM Laboratory, Université de Lorraine. The samples were prepared by mixing 1.5 mL of 50 mM HEPES buffer at pH 7.4 with 250 µL of 10 mM sample solution (hydrolysates: eq. NH₂) and 250 µL of 10 mM nickel(II) sulfate solution in order to have a 1:1 metal-to-ligand molar ratio. The UV-vis spectra were recorded on a Perkin Elmer Lambda1050 UV-vis-NIR spectrophotometer (Perkin Elmer Corp, Massachusetts, USA) in a quartz cuvette with 1.0 cm path length in a range of wavelengths varying from 350 to 800 nm at room temperature (25 °C).

### 2.4.3. CD measurement

The circular dichroism (CD) spectra of the synthetic peptides and the SMPI hydrolysates (whole and ≤ 1 kDa fractions) with and without Ni²⁺ ions were performed at the PhotoNS Platform of the L2CM Laboratory, Université de Lorraine. The CD spectra were recorded from 250 to 800 nm at 0.5 nm intervals using a MOS-450 CD spectrometer (Bio-Logic Science Instruments, France). Solutions of synthetic peptides and hydrolysates were prepared at 10 mM (hydrolysates: mM eq. NH₂) in 50 mM HEPES buffer at pH 7.4 and mixed with 10 mM Ni²⁺ solution to reach 1:1 metal-to-ligand molar ratio. Then, sample solutions (with and without Ni²⁺ ions) were transferred into a 1 cm-pathlength quartz cuvette for analysis at 25 °C as reported in Csire et al. [25].

## 2.5. Statistical analysis

To evaluate the effect of enzymatic treatments, 1 kDa ultrafiltration process and the AA composition and sequences on the Ni²⁺-binding ability of synthetic peptides and SMPI hydrolysates (whole and ≤ 1 kDa fractions), results were statistically analyzed by one-way analysis of variance (ANOVA). All the measurements of Ni²⁺-IMAC results were triplicated for synthetic peptides and SMPI hydrolysates and data were presented as means ± standard deviation. Differences between means of samples were determined by Fisher's multiple range test where differences at *p* < 0.05 were considered significant. Statistical calculations were conducted using the software Statgraphics Centurion XVI version 16.1.03 (Statgraphics Technologies Inc., Virginia, USA).

# 3. Results and discussion

## 3.1. Investigation of Ni²⁺-binding peptides in Ni²⁺-IMAC

### 3.1.1. Retention time of synthetic peptides

In order to study the ability of the different synthetic peptides to bind to Ni²⁺ ions, samples were initially investigated on Ni²⁺-IMAC. This method, based on peptides-Ni²⁺ interaction, provides several advantages due to its high-binding capacity and high recovery. The Ni²⁺-binding peptides were eluted using imidazole gradient (0–300 mM, in 60 min). The Fig. 2 shows the retention time (RT) of the peptides in the column (the red dash line indicates the beginning of imidazole gradient from 0 to 300 mM); the higher the RT (min) or IMC (mM of imidazole), the stronger the Ni²⁺-binding ability of the investigated peptide. The Pep11.HHHHHH, well-known for its strong Ni²⁺-binding ability in IMAC, was used as positive control. It exhibits the highest RT (42.6 min; IMC: 187.9 mM, fraction 2: RT > 5 min), which is associated with its His residues as previously reported [18]. Regarding the sunflower peptides, they were theoretically classified into three groups (see section 2.1): group (I) His or Trp-containing peptides without proline, which is divided into two subgroups IA (His-containing peptides, Pep1-3) and IB (Trp-containing peptide, Pep4); group (II) His-containing peptides with proline (Pep5-8), and group (III) peptides without His nor Trp (Pep9-10). In agreement with the theoretical classification, both His-containing peptides groups IA and (II) without/with proline, respectively 1.WLHNDGN, 2.WLHNDGNTE, 3.FATCIARH, 5.AIPTGTAH, 6.KFPILEH, 7.KFPILEHL and 8.KFPILEHLR exhibit the highest RT with no significant statistical differences (*p* ≥ 0.05). Indeed, these peptides are observed in the elution phase (RT > 5 min; IMC: 22.2–25.2 mM, fraction F2) where imidazole is required for peptide elution. Our results agree with the hard and soft acid and base theory (the so-called HSAB theory), which states that to form a coordination bound between two atoms, one atom behaves like a Lewis base (i.e., imidazole group of histidine) while the other one behaves like a Lewis acid (i.e., Ni²⁺). Hard acids interact more strongly with hard bases than with soft bases, so the most stable complexes are those with hard-hard and soft-soft interactions. The observed interaction between histidine residue and Ni²⁺ ions is governed by both borderline Lewis acid and base groups which confers also the strongest interaction (like binds strongly with like) [26]. Meanwhile, group (III) of peptides, those without His (9.GNPQ, 10.LLAF), were eluted before applying the imidazole gradient (RT < 5 min, fraction F1). In comparison to peptide Pep9.GNPQ and Pep10.LLAF, which have no affinity for Ni²⁺ ions, the Pep4.WVFSFKTN (group IB) has a slightly higher affinity (*p* < 0.05, RT: 4.19 min, fraction F1). Note that, the presence of a Trp residue in the sequence of Pep4.WVFSFKTN, may induce only a weak interaction with Ni²⁺. However, Trp with Cys and His have been proposed by Porath as a specific triad in the IMAC technique as primary electron-pair donors for protein retention [27]. Yet, the presence of a Trp residue is not enough to exhibit a good Ni²⁺-binding ability compared to His residue that has the most significant role in Ni²⁺-binding ability according to the HSAB theory and as reported for sunflower peptides in this study. Besides, the Ni²⁺-IMAC results suggest that
<figure><img src="image_4.png" /><figcaption>**Fig. 2.** Retention times (min) of pure peptides studied with HisTrap X-Ni²⁺ IMAC. Values are expressed as the mean ± standard deviation of three independent determinations. Different letters indicate significant differences (*p* < 0.05).</figcaption></figure>

the affinity of peptides for immobilized Ni²⁺ ions is rather governed by the presence of His residue than by its position within their sequence.

### 3.1.2. Separation of Ni²⁺-binding peptides present in hydrolysate

Similar to synthetic peptides, the Ni²⁺-binding ability of SMPI hydrolysates was investigated using Ni²⁺-IMAC. The Ni²⁺-IMAC chromatograms (Fig. 3A–D) show the injection phase (0–5 min) and elution phase (fraction 2: 5–65 min) of whole hydrolysates and their corresponding ≤1 kDa fractions (the red dash line indicates the beginning of imidazole gradient from 0 to 300 mM). Upon elution phase using an

imidazole gradient (0–300 mM, 60 min from 5 to 65 min RT), Ni²⁺-binding peptides bound onto Ni²⁺-IMAC phase were eluted in one peak, i.e., only one F2 fraction. For all the four studied hydrolysates, a Ni²⁺-binding peptides fraction is observed (F2). Thus, SMPI hydrolysates contain metal-binding peptides. We hypothesize that these potential metal-binding peptides may contain histidine, tryptophane or cysteine (three AA residues called Porath's triad [27]), as well as aspartic and glutamic acids, AA well-known for their transition metal ions binding properties due to their imidazole, indole, thiol and carboxylate groups present on their side chain, respectively [7] [18]. Indeed, at

<figure><img src="image_5.png" /><figcaption>**Fig. 3.** Study of peptide-Ni²⁺ interaction SMPI hydrolysates using immobilized metal ion affinity chromatography (IMAC). (A-D) The isolation of Ni²⁺-binding peptides of SMPI hydrolysates. (E) Retention times (RT) of SMPI hydrolysates studied with HisTrap X-Ni²⁺ IMAC. The unbounding peptides (F1) were eluted by 5 min of equilibrating buffer PBS 1X pH 7.4. Elution profile of Ni²⁺-binding peptides (F2) was obtained after imidazole gradient (5-65 min, 0-300 mM). Values are expressed as the mean ± standard deviation of three independent determinations. Different letters indicate significant differences (*p* < 0.05). SMPI: sunflower meal protein isolate; Prot: Protamex® hydrolysate; Prot+Flav: Protamex® followed by Flavourzyme®; ≤ 1kDa: 1 kDa ultrafiltrated.</figcaption></figure>
physiological pH (7.4), imidazole group of His and COOH group of Asp and Glu are deprotonated, since their side chain group pK values are 6.0, 3.9 and 4.2, respectively, which enable to bind to Ni²⁺ ions [28]. Note that among the hydrolysates, the retention time (RT) of the Ni²⁺-chelating peptides fractions ranged between 9.7 and 10.1 min (IMC: 23.8–25.5 mM), which means that the mechanism of peptide-Ni²⁺ interaction in Ni²⁺-IMAC column may be similar in the four hydrolysates and their difference in chelation capacity may rather be explained by the concentration of chelating peptides, which is related with the absorbance intensity (Fig. 3A–D). Regarding the intensity, the result suggests that Prot hydrolysate contains the highest amount of Ni²⁺-binding peptides (F2) followed by Prot+Flav, ≤ 1 kDa Prot and ≤ 1 kDa Prot+Flav hydrolysates.

### 3.1.3. Ni²⁺-binding peptides identified in hydrolysates by LC-MS/MS

The fractions F1 (containing the unbound peptides) and F2 (enriched in Ni²⁺-binding peptides) collected from previous chromatographic experiments were subjected to LC-MS/MS analysis in order to identify the peptide sequences present in SMPI hydrolysates. The peptide sequences identified in the enriched Ni²⁺-binding peptides fractions (F2) are shown in Table 2. Interestingly, all the peptide sequences identified in F2 fractions, contain one His (except for GSRWVSF which contains one Trp residue), which could play a pivotal role in the Ni²⁺-binding ability of peptides present in hydrolysates due to its imidazole (or indole group for GSRWVSF) (Table 2, H or W residues highlighted in bold). In comparison, among the 175 peptides identified in the unbound peptides fractions F1, only 1 peptide contains His (DFIRH) and 5 peptides contain Trp (i.e., EIWDANDQ, EWVAF, REEQEWE, SLWN, WTVN; Table S4). This finding agrees with the low Ni²⁺-binding ability of the synthetic peptide Pep4.WVFSKTN (Trp-containing peptide), which was also eluted before applying the imidazole gradient (RT < 5 min).

The Table 2 reveals a total number of 22, 10, 22 and 9 Ni²⁺-binding peptides identified in Prot, Prot+Flav, ≤ 1 kDa Prot and ≤ 1 kDa Prot+Flav, respectively. From Table 2, Fig. 4A illustrates the effect of the

sequential enzymatic treatment using Flav on the generation of Ni²⁺-binding peptides. Upon the Flav sequential hydrolysis, 15 Ni²⁺-binding peptides already produced in Prot hydrolysate are hydrolysed (thus disappearing; represented by a cross (†) in Table 2), whilst 3 additional peptides are generated (i.e., GHLQPN, HNDGNTELVVVVF and SFLAG-GISSEHIQGQQQ, marked by an asterisk (*) in Table 2). Moreover, Fig. 4B shows the effect of ≤ 1 kDa ultrafiltration on the appearance of peptides that might not be identified in the whole hydrolysates due to their concentration below the limit of detection of MS/MS analysis. After ultrafiltration, no additional peptides were identified in ≤ 1 kDa Prot fraction. Besides, two big-sized peptides (HNDGNTELVVVVF and SFLAG-GISSEHIQGQQQ, with MW > 1.4 kDa) disappeared upon ultrafiltration in ≤ 1 kDa Prot+Flav (which is coherent with the MWCO of the UF process), while an additional peptide was identified (SEVFH) probably related by a better relative detection in the ultrafiltrated hydrolysate (Table 2).

In addition to the presence of His residue in the peptide sequence, the size and the AA composition of Ni²⁺-binding peptides constituting the fraction F2 plays a pivotal role on the Ni²⁺-binding ability. From Table 2 data, the Fig. 4C displays the fraction of Ni²⁺-binding peptides (%) calculated for different peptide lengths (number of AA residues). The length of the identified Ni²⁺-binding peptides ranges between 3 and 17 AA residues. For whole hydrolysates and ≤ 1 kDa fractions, the majority of peptide fraction comprises between 5 and 7 AA (> 50 % peptides). Besides, both Prot hydrolysates (whole and ≤ 1 kDa fraction) were the only ones, which present small-sized Ni²⁺-binding peptides constituted between 3 and 4 AA in the fraction F2. This suggests that although the sequential hydrolysate (Prot+Flav) exhibits higher overall production of small-sized peptides compared to Prot hydrolysate (higher degree of hydrolysis, Table S1), sequential treatment does not favor the production of small-sized Ni²⁺-binding peptides (3–4 AA residues).

The AA composition of Ni²⁺-binding peptides (fraction F2) was analyzed by the free-accessed ProtParam tool (https://web.expsasy.org/protparam/) (Fig. 4D). It evidences that all the hydrolysates

<table><caption>**Table 2**
Ni²⁺-binding peptides identified from the F2 (elution phase) SMPI hydrolysates using IMAC-Ni²⁺ separation followed by LC-MS/MS analysis.</caption><thead><tr><th rowspan="2">Sequence</th><th colspan="4">Peptides identified in F2 fraction</th><th rowspan="2">Length (AA)</th><th rowspan="2">Mass (Da)</th><th rowspan="2">Score</th><th rowspan="2">Entry</th></tr><tr><th>Prot</th><th>Prot + Flav</th><th>≤ 1 kDa Prot</th><th>≤ 1 kDa Prot + Flav</th></tr></thead><tbody><tr><td>AHNLL†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>5</td><td>566.32</td><td>1.94</td><td></td></tr><tr><td>AHNLLY†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>6</td><td>729.38</td><td>4.12</td><td></td></tr><tr><td>AIQSPHWT†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>8</td><td>938.46</td><td>5.76</td><td>A0A251TKL9</td></tr><tr><td>AIQSPHWTIN</td><td>x</td><td>x</td><td>n.i.</td><td>n.i.</td><td>10</td><td>1165.59</td><td>6.72</td><td></td></tr><tr><td>ALFSPH†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>6</td><td>670.34</td><td>9.64</td><td>A0A251UYX1</td></tr><tr><td>AVLAPH†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>6</td><td>606.35</td><td>9.64</td><td>A0A251T566</td></tr><tr><td>DFIRH</td><td>x</td><td>x</td><td>x</td><td>x</td><td>5</td><td>686.35</td><td>16.21</td><td>A0A251TKL9</td></tr><tr><td>FSPH†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>4</td><td>486.22</td><td>8.08</td><td>A0A251UYX1</td></tr><tr><td>GAAHW†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>5</td><td>540.24</td><td>1.94</td><td>A0A251TUS1</td></tr><tr><td>GAVHW†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>5</td><td>568.28</td><td>6.05</td><td>A0A251T566</td></tr><tr><td>GHLQPN*</td><td>n.i.</td><td>x</td><td>n.i.</td><td>x</td><td>6</td><td>664.33</td><td>9.64</td><td>A0A251UYX1</td></tr><tr><td>GIVSPH</td><td>x</td><td>x</td><td>x</td><td>x</td><td>6</td><td>608.33</td><td>9.64</td><td>A0A251TUS1</td></tr><tr><td>GSRWVSF†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>7</td><td>837.41</td><td>2.89</td><td></td></tr><tr><td>GVDIFIRH</td><td>x</td><td>x</td><td>x</td><td>x</td><td>7</td><td>842.44</td><td>17.09</td><td>A0A251TKL9</td></tr><tr><td>GVDFIRHR</td><td>x</td><td>x</td><td>x</td><td>x</td><td>8</td><td>998.54</td><td>3.60</td><td></td></tr><tr><td>HAW†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>3</td><td>412.19</td><td>3.98</td><td>A0A251VMR6</td></tr><tr><td>HNDGNTELVVVVF*</td><td>n.i.</td><td>x</td><td>n.i.</td><td>n.i.</td><td>13</td><td>1441.72</td><td>1.46</td><td>A0A251TKL9</td></tr><tr><td>LLHVT†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>5</td><td>581.35</td><td>5.30</td><td>A0A9K3NGE6</td></tr><tr><td>SEVFH</td><td>n.i.</td><td>n.i.</td><td>n.i.</td><td>x</td><td>5</td><td>617.28</td><td>1.94</td><td>A0A9K3HVJ0</td></tr><tr><td>SFLAGGISSEHIQGQQQ*</td><td>n.i.</td><td>x</td><td>n.i.</td><td>n.i.</td><td>17</td><td>1785.86</td><td>0.35</td><td>A0A251UYX1</td></tr><tr><td>SHTVVY†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>6</td><td>704.35</td><td>20.22</td><td></td></tr><tr><td>SPH†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>3</td><td>339.15</td><td>3.98</td><td></td></tr><tr><td>SPHWT</td><td>x</td><td>x</td><td>x</td><td>x</td><td>5</td><td>626.28</td><td>1.65</td><td>A0A251TKL9</td></tr><tr><td>SPHWTIN</td><td>x</td><td>x</td><td>x</td><td>x</td><td>7</td><td>853.41</td><td>11.11</td><td></td></tr><tr><td>VSPH†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>4</td><td>438.22</td><td>8.08</td><td>A0A251TUS1</td></tr><tr><td>WLH†</td><td>x</td><td>n.i.</td><td>x</td><td>n.i.</td><td>3</td><td>454.23</td><td>3.98</td><td>A0A251TKL9</td></tr><tr><td>Total peptides</td><td>22</td><td>10</td><td>21</td><td>8</td><td></td><td></td><td></td><td></td></tr></tbody></table>

SMPI: sunflower meal protein isolate; Prot: Protamex® hydrolysate; Prot+Flav: Protamex® followed by Flavourzyme®; ≤ 1 kDa: 1 kDa ultrafiltrated. n.i.: peptide not identified; x: peptide identified in the F2 fraction; †: 15 peptides disappeared upon sequential treatment; *: 3 peptides produced upon sequential treatment; score: measure of how well the observed MS/MS spectrum matches to the stated peptide.
<figure><img src="image_6.png" /><figcaption>**Fig. 4.** Ni²⁺-binding peptides present in SMPI hydrolysates identified by the LC-MS/MS. (A-B) Effect of sequential enzymatic treatment and 1 kDa ultrafiltration on the number of Ni²⁺-binding peptides identified in F2 fractions. (C) Peptide length profile of identified peptides. (D) AA frequency of Ni²⁺-binding peptides identified. AA: amino acids; Prot: Protamex® hydrolysate; Prot+Flav: Protamex® followed by Flavourzyme®; ≤ 1kDa: 1 kDa ultrafiltrated.</figcaption></figure>

contain His residue as the major amino acid in their sequences (100 %), histidine being the main AA residue, responsible of their Ni²⁺-binding properties. Moreover, Fig. 5A–L illustrates the sequenced mass spectra of the most interesting sequences of Ni²⁺-binding peptides from SMPI hydrolysates loaded into the MaxQuant Viewer. The mass spectra of sequences with His residue at third position from N-terminus, second and first position from both termini are shown in Fig. 5A–D, Fig. 5E–H, and Fig. 5I–L, respectively. In our study, due to the presence of His residue as well as its position within the sequence, the identified sequences could be endowed with the highest metal-binding properties. Indeed, the Ni²⁺-binding peptides found in SMPI hydrolysates with His residue at first, second and third position from the N-terminus could possess suitable Ni²⁺-binding ability and stability in the formed peptide-M2⁺ complex as previously reported [29]. The Prot hydrolysates (whole and ≤ 1 kDa fraction) shows high peptide sequences with His residue placed near the N-terminus while there are only four in the case of Prot+Flav hydrolysates (Table 2). Interestingly, these hydrolysates contain LLHVT and WLH sequences (Fig. 5A–B), which are the most interesting ones due to the presence of their so-called “amino terminal CuNi(II)-binding (ATCUN) motif. This ATCUN motif is structurally characterized by (i) a free N-terminal amino nitrogen (NH₂) of the first residue, (ii) two deprotonated amide nitrogens of the peptide backbone, and (iii) an imidazole nitrogen of the third His residue from N-terminus [30] (Fig. 1L). This ATCUN motif has been highly used to design Ni²⁺-binding peptides due to its high Ni²⁺-binding ability, specificity and stability [31–34]. Additionally, apart from this ATCUN motif, several sequences identified by mass analysis contain the His residue at the second position from N-terminus (i.e., AHNLLY, AHNLL, SHTVVY for Prot hydrolysates; GHLQPN for Prot+Flav hydrolysates, Table 2), which could also offer interesting Ni²⁺-binding ability. Indeed, Conato et al. [35] reported that a Ni²⁺-chelating peptide with such a structural feature (i.e., His residue in second position, GHK) possess strong Ni²⁺-binding affinity. Besides, for peptides with His residue at first and second position from C-terminus (i.e. GVDFIRH, GAAHW), both terminal amino and imidazole side chains can take part in the Ni²⁺ complexation;

however, these sites could act as independent metal binding sites when they are far apart in the sequence [29].

On the other hand, there are many Ni²⁺-binding peptide sequences identified that, in addition to the His residue, contain proline as well (Table 2). Yet, the Ni²⁺-binding properties of these particular identified peptides might be lower compared to others. Indeed, according to the literature, proline could usually work as a break-point for amide coordination since it cannot be deprotonated, affecting the stability of the formed peptide-metal complex [16]. Former authors demonstrated that the effect of proline on the formation of peptide-Ni²⁺ complex depends on its position from N-terminus, the highest effect of proline on the complexation can be found when it precedes directly the His residue [16]. Some of the Ni²⁺-binding peptides identified in Table 2 (e.g. VSPH, SPHWT) may exhibit this behaviour. Similarly to the synthetic peptides, hydrolysates investigated by Ni²⁺-IMAC do not show potential differences between the Ni²⁺-binding ability of His-containing peptides when proline is present, therefore, other technologies involving immobilized Ni²⁺ ions are required.

Overall, in our study, data obtained from Ni²⁺-IMAC separation (fraction F2) followed by peptide sequence identification by LC-MS/MS analysis are used to better understand the metal binding behaviour of peptides present in whole SMPI hydrolysates and their ≤ 1 kDa fractions and are complemented by other techniques such as switchSENSE® technology, UV-visible and circular dichroism spectroscopies. While the investigation by these techniques provides information on metal-binding constant rates, coordination geometry as well as potential binding sites of specific peptides, understanding the peptide sequence composition of the Ni²⁺-binding fractions in IMAC (fraction F2) can be essential to decipher the Ni²⁺-binding ability and complexation mechanisms involved in a mixture of peptides such as SMPI hydrolysates.
<figure><img src="image_7.png" /><figcaption>Fig. 5. Sequenced mass spectra of twelve different Ni²⁺-binding peptides structures obtained from SMPI hydrolysates and processes by MaxQuant software. ATCUN motif-containing peptides identified: (A) LLHVT, (B) WLH, (C) SPH and (D) SPHWTIN; His-containing peptide with His residue placed at second position from N and C-termini: (E) AHNLLY, (F) SHTVVY, (G) GAVHW, (H) GVDFIRHR; His-containing peptide with His residue placed at first position from N and C-termini: (I) HAW, (J) HNDGNTELVVVVF, (K) DFIRH, (L) SEVFH.</figcaption></figure>
## 3.2. Metal-peptides interaction

### 3.2.1. Affinity parameter determination by switchSENSE® technology

#### 3.2.1.1. Static measurement.
Ni²⁺-binding ability of synthetic peptides and SMPI hydrolysates was also evaluated by real-time molecular interaction analysis using the switchSENSE® technology. Similarly to Ni²⁺-IMAC, this technology studies the interaction between peptides and immobilized Ni²⁺ ions, allowing to unveil potential differences of

the degree of Ni²⁺-binding affinity of sunflower peptides (synthetic peptides or hydrolysates).

The kinetic study of the interaction between synthetic peptides and Ni²⁺ reveals that peptides are bound onto the immobilized Ni²⁺ ions with different binding rate ($k_{on}$) (Fig. S3A-K), while no dissociation phase ($k_{off}$) could be observed within the time of measurement; thus, the equilibrium dissociation constant $K_D$ values ($K_D = k_{off}/k_{on}$) could not be determined as previously reported [19]. Instead, the global fitting by single exponential functions to the raw data in the association phase

<figure><img src="image_8.png" /><figcaption>**Fig. 6.** Study of the interaction of synthetic peptides and SMPI hydrolysates with nickel ions using SwitchSENSE® technology. SwitchSENSE® measurements in static mode: (A-B) average observed rate constant ($k_{obs}$) of synthetic peptides and SMPI obtained from kinetic study. (C-D) Changes in the local fluorophore environment (DF) of free-nickel references spots and samples spots for synthetic peptides and SMPI hydrolysates. SwitchSENSE® measurements in dynamic mode: (E-F) changes in the switching speed (DDR) of nanolevers upon peptides binding into Ni²⁺ ions for synthetic peptides and SMPI hydrolysates. Values are expressed as the mean ± standard deviation. Different letters indicate significant differences ($p < 0.05$). SMPI: sunflower meal protein isolate; Prot: Protamex® hydrolysate; Prot+Flav: Protamex® followed by Flavourzyme®; $\le 1$ kDa: 1 kDa ultrafiltrated.</figcaption></figure>
(solid lines in Fig. S3A-K) allowed to determine the observed rate constant ($k_{obs}$) (Fig. 6A). The obtained $k_{obs}$ values provide information about how fast the binding reaction between peptide and metal ion occurs and thus allow the comparison of synthetic peptides with each other and also hydrolysates at a defined concentration. Upon the $Ni^{2+}$ binding, several behaviors are observed as described hereafter. First and as expected, Pep11.HHHHH exhibits the highest $Ni^{2+}$-binding association rate ($k_{obs} = 6.72 \times 10^{-2} s^{-1}$) as positive control. Secondly, the Pep1.WLHNDGN, 2.WLHNDGNT, 3.FATCIARH and 4.WVSFKTN (initially classified as the group I according to HSAB theory) present the highest association rate among synthetic peptides ($3.04–3.54 \times 10^{-2} s^{-1}$), with no significant statistical differences ($p \ge 0.05$). Thirdly, the Pep5.AIPTGTAH, Pep6.KFPILEH, 7.KFPILEHL and 8.KFPILEHLR binds to $Ni^{2+}$ ions with lower binding rate than Pep1-4 ($p < 0.05$). Note that these latter group II peptides bind $Ni^{2+}$ ions with similar binding rate ($k_{obs} = 1.59–2.45 \times 10^{-2} s^{-1}$) which is the lowest association rate for His-containing peptides. Finally, Pep9.GNPQ and 10.LLAF (group III peptides), which do not contain any His nor Trp residue in their sequence, show the lowest $k_{obs}$ values, demonstrating they have hardly any affinity for immobilized $Ni^{2+}$ ions.

Concerning the hydrolysates (whole and ultrafiltrated), the kinetic study of the interaction between peptides contained in SMPI hydrolysates and immobilized $Ni^{2+}$ ions notably shows some variation in fluorescence during the binding process (association phase, Fig. S4A-D) confirming the presence of $Ni^{2+}$-binding peptides in all the SMPI hydrolysates and their respective $\le 1$ kDa fractions. Similarly to the synthetic peptides, the global fitting of the raw data in the association phase of the investigated hydrolysates allowed to determine their observed rate constant ($k_{obs}$, Fig. 6B). The $Ni^{2+}$-binding peptides present in Prot hydrolysate bind to $Ni^{2+}$ ions much faster ($31.28 \times 10^{-2} s^{-1}$, $p < 0.05$) than those present in Prot+Flav one ($k_{obs} = 0.15 \times 10^{-2} s^{-1}$), whereas among their respective $\le 1$ kDa fractions, the $k_{obs}$ values decrease and are not statistically different ($p > 0.05$). This observation clearly shows that the high $Ni^{2+}$-binding properties of Prot hydrolysate is strongly associated with its higher concentration of $Ni^{2+}$-binding peptides compared to other hydrolysates, which agrees with the above-mentioned results found in $Ni^{2+}$-IMAC.

Besides, for a more accurate assessment of the $Ni^{2+}$ binding ability of the peptides and hydrolysates, relative changes in fluorescence ($\Delta F$) were measured on sample spots (i.e., spots functionalized with $NTA-Ni^{2+}$) compared to reference spots (i.e., REF: spots in the absence of $NTA-Ni^{2+}$) upon injection of the peptide or hydrolysates (Fig. 6C-D). Overall, all the studied peptides present a shift of $\Delta F$ when compared with REF. Nevertheless, the His or Trp-containing peptides of group I and II (Pep1.WLHNDGN, 2.WLHNDGNT, 3.FATCIARH, Pep4.WVSFKTN, 5.AIPTGTAH, 6.KFPILEH, 7.KFPILEHL and 8.KFPILEHLR), and the positive control (Pep11.HHHHH) evidence much higher shift of $\Delta F$ compared with REF values than peptides without His or Trp (group III peptides: Pep9.GNPQ and 10.LLAF). This validates the affinity of His or Trp-containing peptide (group I and II) for $Ni^{2+}$ ions compared to peptides without these specific AA. Regarding hydrolysates (whole or ultrafiltrated), all the samples show significant differences compared with the REF ($p < 0.05$), confirming the presence of $Ni^{2+}$-binding peptides.

### 3.2.1.2. Dynamic measurement.
Dynamic mode measurements enable to analyze the $Ni^{2+}$-binding ability of peptides based on a decrease of the switching speed of nanolever upon the $Ni^{2+}$ binding through peptides. The measurements were carried out, between the association and dissociation phase of the static mode while applying a current voltage. In this dynamic mode, the switching speed decreases upon the peptide binding onto $Ni^{2+}$ since the nanolevers becomes heavier and its hydrodynamic friction is increased.

As shown in Fig. 6E, Pep1.WLHNDGN, 2.WLHNDGNT and 3.FATCIARH exhibit the highest change of switching speed ($\Delta$DR compared to

REF) on the motion of nanolevers upon the $Ni^{2+}$ binding, followed by Pep4-10 (4.WVSFKTN, 5.AIPTGTAH, 6.KFPILEH, 7.KFPILEHL and 8.KFPILEHLR, 9.GNPQ and 10.LLAF) with the lowest variation, associated with their low affinity for $Ni^{2+}$ ions showed in static mode (the lowest $k_{obs}$ values). In addition to their His residue, Pep1.WLHNDGN, 2.WLHNDGNT and 3.FATCIARH are characterized by the presence of Trp or Cys residues, respectively. These three amino acids (His, Cys and Trp,) constitutes the so-called Porath's triad [27]; due to their respective presence of imidazole (His), thiol (Cys) and indole (Trp) groups, they could therefore also be an anchoring sites for transition metal ions [36].

Regarding SMPI hydrolysates, Fig. 6F reveals that, compared to other hydrolysates, the $Ni^{2+}$-binding peptides present in Prot hydrolysate show some effect on the motion of nanolevers, since the dynamic response ($\Delta$DR) for sample spot compared to REF is less than 100 % upon the $Ni^{2+}$ binding (97 %, $p < 0.05$). Indeed, Prot hydrolysate, characterized by lower degree of hydrolysis (DH) than Prot+Flav hydrolysate (Table S1), may contain higher concentration of bigger-sized $Ni^{2+}$-binding peptides causing higher hydrodynamic friction. In agreement with static mode measurement, Prot hydrolysates shows the most interesting $Ni^{2+}$-binding ability, whereas sequential Prot + Flav hydrolysate and the $\le 1$ kDa fractions evidence a negligible effect ($p > 0.05$) on the motion of nanolevers upon the binding which is related to both their low $k_{obs}$ values observed and also the small-sized $Ni^{2+}$-binding peptides identified.

Taken together, all the findings of switchSENSE® technology on synthetic peptides clearly show that group I peptides (Pep1.WLHNDGN, 2.WLHNDGNT, 3.FATCIARH and 4.WVSFKTN, His or Trp-containing peptides without Pro residue) have stronger interaction with the $Ni^{2+}$-binding sites than group II peptides (Pep5.AIPTGTAH, 6.KFPILEH, 7.KFPILEHL and 8.KFPILEHLR, His-containing peptides with Pro residue) and group III (9.GNPQ and 10.LLAF, peptides without His or Trp residue). Although group I and II peptides contain the same number of His $Ni^{2+}$-binding ligand (one His residue in their sequence), the interaction towards $Ni^{2+}$ ion is lower in this last group (group II containing His and Pro residue). To explain this observation, one can mention the position of the effective anchoring site for metal ion coordination, which is located at the C-terminus in group II peptides. They can therefore only form a {COO$^-$, $N_{His}$} chelate ring or {NH$_2$, $N_{His}$} macrochelate, thermodynamically less stable than those formed by the group I peptide {NH$_2$, $N_{His}$} where His is at the N-terminal end. The Pep.3 from group I could be an exception with its C-term His. Here, in addition to the 7-membered chelate ring formation at the C-term, the stability of this $Ni^{2+}$ complex is enhanced by the presence of Cys in the sequence. Indeed, it has also been demonstrated that the thiol group can be an efficient anchor and coordinating group for $Ni^{2+}$ ion providing additional stability to complex [37]. As in all cases, the relative position of AA residues with coordinating side chains has a crucial impact on the binding mode, thus on stability. In Pep3.FATCIARH, Cys and His are far apart to exhibit the cooperative effect in amide deprotonation, but Cys acts as a supplementary coordinating group, adding extra stability relative to the group II peptides. Another point to explain the difference in chelating behaviour between Pep.3 and group II, is the {NH$_2$, $N_{His}$} macrochelate formation, which can be sterically blocked by the presence of rigid Proline.

Additionally, but a lesser extend in the case of ternary complex formation between $NTA-Ni^{2+}$ and peptides, it was hypothesized that the presence of Pro residue in Pep5 to 8 (Pep5.AIPTGTAH, 6.KFPILEH, 7.KFPILEHL and 8.KFPILEHLR) could attenuate their $Ni^{2+}$-binding mechanism. Since Pro is a secondary amine AA, which after its involvement in peptide bond cannot be deprotonated, and therefore cannot take part in metal binding as observed for the ATCUN-motif [16].

These findings suggest that, in our study, the $Ni^{2+}$-binding ability and the strength of binding of the peptides to $Ni^{2+}$ ions might not depend only on the number of His residues, but also to their position within the peptide sequence, which influences their structural conformation. Indeed, the arrangement of His residue, the adjacent amino acids and
the type of amino acid at the termini of Ni²⁺-binding peptides play a key role on Ni²⁺ complexation.

Finally, interesting conclusions from the switchSENSE® data of synthetic peptides can also be drawn in SMPI hydrolysates: (i) the Ni²⁺-binding ability of peptide sequences identified in sunflower hydrolysates (Table 2), which could be classified in peptides from the group I (His or Trp-containing peptides without proline; e.g. AHNLL, LLHVT, WLH) and group II (His-containing peptides with proline; e.g. AIQSPHWTIN, GHLQPN, SPHWT), would vary depending on the presence of His and Pro residue as well as on their position within the peptide sequence. Besides, the ATCUN motif-containing peptides identified in Prot hydrolysates, particularly LLHVT and WLH (present in Pep1.WLHNDGN, 2.WLHNDGNTE) as well as all the other His-containing peptides without proline present in SMPI hydrolysates (Table 2), could have the best Ni²⁺-binding properties among the identified Ni²⁺-binding peptides in SMPI hydrolysates.

### 3.2.2. UV-visible spectroscopy and CD measurements

UV-visible spectroscopy (UV-vis) and circular dichroism (CD) were used to study the coordination characteristics of the resulting complex of synthetic peptides or SMPI hydrolysates with Ni²⁺ ions in solution at physiological pH (7.4). The coordination modes of peptide-Ni²⁺ complexes were investigated by comparing the absorption/CD band maxima of the peptides alone with those of the corresponding peptide-Ni²⁺ complex using the same concentration. Thanks to UV-vis and CD measurements, different electronic transitions commonly found in metal coordination complexes can be observed. Among these specific electronic transitions, the d-d transition referring to the electronic transition between different d-orbitals of the metal ion (i.e., Ni²⁺) appears as relatively weak absorptions. In comparison, the charge transfer (CT) transition which involves an electronic transition from the molecular orbital of the peptide ligand to the metal ion is characterized by stronger intensities and shorter wavelength in the absorption spectrum than the d-d transition. As example, Fig. 7A-F and Fig. 8A-H show the UV-vis absorption and CD behaviors of peptide-Ni²⁺ complexes for three

<figure><img src="image_9.png" /><figcaption>Fig. 7. UV-visible and CD measurements of the peptide-Ni²⁺ interaction of pure peptides. (A-B) Pep2.WLHNDTE; (C-D) Pep3.FATCIARH; (E-F) Pep11.HHHHHH.</figcaption></figure>
<figure><img src="image_10.png" /><figcaption>Fig. 8. UV-visible and CD measurements of the SMPI hydrolysates (A-B) Prot; (C-D) Prot+Flav; (E-F) ≤ 1 kDa Prot; (G-H) ≤ 1 kDa Prot+Flav. SMPI: sunflower meal protein isolate; Prot: Protamex® hydrolysate; Prot+Flav: Protamex® followed by Flavourzyme®; ≤ 1kDa: 1 kDa ultrafiltrated. Horizontal arrow indicates shift in wavelength after formation of peptide-Ni²⁺ complex.</figcaption></figure>
synthetic peptides (i.e., 2.WLHNDGNT, 3.FATCIARH and 11. HHHHHH) and SMPI hydrolysates, respectively. From such spectra, spectroscopic data (absorption maxima and sign of + or – Cotton effect on CD) were summarized in Table 3 for all investigated peptides and hydrolysates.

**Peptides.** Among the synthetic peptides, only the Pep11.HHHHHH (positive control) and the His or Trp-containing peptides belonging to the group IA (Pep1.WLHNDGN not shown, 2.WLHNDGNT, and 3. FATCIARH) evidence measurable changes in the absorption behaviour of peptide upon their Ni²⁺ complexation. They are characterized by a redshifted d-d band at around 420 nm relative to the band of Ni²⁺ alone found in water solution at 390 nm (blue line; Fig. 7). Indeed, these peptides exhibit the most interesting Ni²⁺-binding ability as previously observed in Ni²⁺-IMAC and switchSENSE® techniques. Yet, for the group II peptides (Pep5.AIPTGTAH, 6.KFPILEH, 7.KFPILEHL and 8. KFPILEHLR), no measurable changes in UV-vis/CD spectra were detected at pH 7.4 (data not shown), although the Ni²⁺ ion is complexed. A possible explanation is that under physiological conditions (pH 7.4) only macrochelates {NH₂, Nᵢₘ} are formed for this peptide group, which do not cause significant changes in the UV-vis spectra. Similarly, for peptides lacking histidine and/or cysteine, such as Pep4.WVSFKTN and group III peptides (9.GNPQ and 10.LLAF), no measurable changes were observed in the UV-vis/CD spectra at pH 7.4. This is consistent with the fact that the formation of square planar nickel(II) complexes in peptides with non-coordinating side chains occurs only at alkaline pH, where the terminal amino group and deprotonated amide donor atoms participate in coordination. In particular, these complexes form above pH 8 when metal-ion assisted cooperative deprotonation of two or three amide groups can occur [37].

Pep1.WLHNDGN and Pep2.WLHNDGNT contain the same AA motif (WLHNDGN) with a slight difference in C-terminus in Pep2 (the TE motif). However, the UV-vis and CD data are similar for both peptides, suggesting that these additional amino acids residues (i.e., threonine and glutamic acid) do not have any important effects on the Ni²⁺ coordination mode at pH 7.4 (Table 3). Therefore, in order to study the complex geometry and the binding modes of WLH motif present in Pep1 and 2, only the UV-vis and CD data of Pep2.WLHNDGNT were used.

The analysis of spectroscopic data presented in Table 3 reveals that Pep11.HHHHHH and 2.WLHNDGNT show similar behaviour upon Ni²⁺ complexation at pH 7.4 (λmax = 420 nm, CD bands in the visible range at 420 nm (+) and 485 nm (–), and agree well with earlier results published on hexahistidine-Ni²⁺-complex [38]. This UV-vis/CD behaviour (Fig. 7A–B and Fig. 7E–F) is consistent with a square-planar environment, and strongly supports a 4N-binding mode with {NH₂, 2Nₐₘide, Nᵢₘidazole} donor set in the formed peptide-Ni²⁺ complex. Indeed, due to their binding mode, the N-terminal histidyl peptides Pep1.WLHNDGN and Pep2.WLHNDGNT, characterized by the ATCUN motif within their sequence “Xaa-Yaa-His-Zaa” (Xaa, Yaa and Zaa meaning any AA) are likely to exhibit a high Ni²⁺-binding capacity [37].

Turi et. [39] reported the Ni(II)-complex formation process of His-

containing peptide (NH₂-MKHM-NH₂ and NH₂-GTHS-NH₂ and octa-peptides NH₂-GTHSMKHM-NH₂, NH₂-MKHMGTHS-NH₂) with His residue at third position from N-terminus as Pep2.WLHNDGNT in our study. Authors suggested this 4N {NH₂, 2Nₐₘide, Nᵢₘidazole} coordination mode where imidazole and terminal NH₂ groups are the primary Ni²⁺ binding sites followed by the deprotonation and coordination of two amide nitrogen atoms in a cooperative manner forming a (5,5,6)-membered fused chelates rings at physiological pH. The CD spectral pattern of Pep2.WLHNDGNT investigated in our study (containing the WLH ATCUN motif) is similar to that of [MLH₂] species proposed.

Besides, the CD profile of Pep3.FATCIARH, characterized by its His residue at first position from C-terminus, shows two bands at 280 nm (– Cotton effect) and 337 nm (+ Cotton effect), and also two distinct bands in the visible region at 462 nm (– Cotton effect) and 553 nm (+ Cotton effect), suggesting a different binding mode compared to Pep11 and 2 (Fig. 7D). In the case of this oligopeptide containing only a single histidyl residue, the simultaneous coordination of Ni(II) ion by the N-terminus and the imidazole-N donor atom in C-terminal location provides the possibility of macrochelation. At physiological pH the amide deprotonation can start from both termini forming 3N ({NH₂, 2Nₐₘide} or {Nᵢₘidazole, 2Nₐₘide}) species characterized by this specific CD profile (4N species ({NH₂, 3Nₐₘide} or {Nᵢₘidazole, 3Nₐₘide}) can be formed at higher pH). Since for group II peptides (Pep5.AIPTGTAH, 6.KFPILEH, 7. KFPILEHL and 8.KFPILEHLR) where the presence of proline in the 3rd position from N-term does not allow the successive amide deprotonation, no measurable CD spectra were observed, confirming the hypothesis that for Pep3.FATCIARH this deprotonation can only occur at the N-term. However, we cannot completely exclude the coordination of imidazole because for Pep10.LLAF of group III also no modification was observed on the CD spectrum demonstrating that no amide deprotonation occurs from the N-term.

**Hydrolysates.** As shown in Fig. 8A–H and Table 3, upon the Ni²⁺ complexation through peptides present in SMPI hydrolysates and their ≤1 kDa fractions, the formation of peptide-Ni²⁺ complex results in a very slight blue shift of the d-d transition band centred at 387 nm. Additionally, some variation is more evident in the 600–700 nm region, where the two bands at 652 and 713 nm from Ni²⁺ alone are shifted to a single broad band around 668–688 nm (Fig. 8A, C, E, F). This finding agrees with the observation reported by Matys et. [40], who found two d-d transition bands shifts between 370 and 690 nm upon the formation of CNAKKHPRCGGG-Ni²⁺ complex in the UV-vis absorption spectrum. Our spectral data shows that the formation of peptide-Ni²⁺ complex could occur in a similar manner for all the four studied hydrolysates. These UV-vis profiles can be attributed to a coordination sphere in octahedral geometry (absorbance around 370–390 and 600–700 nm [41] [42]), but in a mixture containing several Ni²⁺ coordinating peptides, it is difficult to identify a single coordination type. The measured spectrum is the addition of different individual spectra probably representing different type of coordination mode at physiological pH. However, CD data analysis (Fig. 8B, D, F, H) can provide more information of

<table><caption>**Table 3**
Spectroscopic data for the peptide-Ni²⁺ complexes at 1:1 metal-to-ligand ratio at pH 7.4.</caption><thead><tr><th rowspan="2">Sample</th><th colspan="3">UV-vis (nm)</th><th colspan="3">CD (nm)</th></tr><tr><th>390</th><th></th><th></th><th></th><th></th><th></th></tr></thead><tbody><tr><td>Ni²⁺</td><td></td><td></td><td>652</td><td>713</td><td></td><td>N.A</td></tr><tr><td>Pep1.WLHNDGN Pep2.WLHNDGNT</td><td>418</td><td></td><td></td><td></td><td>300 (+)</td><td>418 (+)</td><td>485 (–)</td></tr><tr><td>Pep3.FATCIARH</td><td></td><td>426</td><td>545</td><td></td><td>280 (–)</td><td>337 (+)</td><td>462 (–)</td><td>553 (+)</td></tr><tr><td>Pep4–10*</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Pep11.HHHHHH</td><td>420</td><td></td><td>622</td><td></td><td>274 (+)</td><td>420 (+)</td><td>486 (–)</td></tr><tr><td>Prot</td><td>387</td><td></td><td></td><td>678</td><td></td><td></td><td>485 (–)</td></tr><tr><td>Prot+Flav</td><td>385</td><td></td><td></td><td>668</td><td></td><td>420 (+)</td><td>497 (–)</td></tr><tr><td>≤ 1 kDa Prot</td><td>388</td><td></td><td></td><td>688</td><td></td><td></td><td>484 (–)</td></tr><tr><td>≤ 1 kDa Prot+Flav</td><td>388</td><td></td><td></td><td>677</td><td></td><td></td><td>500 (–)</td></tr></tbody></table>

N.A: not applicable.

– not significant changes upon the addition of Ni²⁺ ions.

* Pep4: WVSFKTN, Pep5: AIPTGTAH, Pep6: KFPILEH, Pep7: KFPILEHL, Pep8: KFPILEHLR, Pep9: GNPQ, Pep10: LLAF.
the binding properties in these peptides mixtures. One main band is observed with a negative Cotton effect around 484–500 nm and a weak positive band at 420 nm only for Prot+Flav hydrolysate. Comparing these spectra with the CD profiles of synthetic Pep11.HHHHHH, Pep2. WLHNDGNT and Pep3.FATCIARH, we can conclude that in all hy- drolysate the presence of {NH₂, 2N<sub>amide</sub>, N<sub>imidazole</sub>} coordination observed for Pep11.HHHHHH and Pep2.WLHNDGNT, as well as the Pep3.FATCIARH-type coordination mode are present but this latter is more predominant in Prot hydrolysate as the observed maximum is slightly blue shifted (Table 3). This hypothesis can be supported by the identified Ni²⁺-binding peptides found in the F2 (elution phase) of SMPI hydrolysates (Ni²⁺-IMAC separation followed by LC-MS/MS analysis, Table 2). Based on these data, the Prot hydrolysate contains more pep- tide having the sequence with histidine at the C-terminus, such as GAAHW, GAVHW or AIQSPHWT which are disappeared upon sequen- tial treatment.

In our study, for the first time to the author's knowledge, we have successfully identified the structures of Ni²⁺-binding peptides present in sunflower meal hydrolysate using Ni²⁺-IMAC-LC-MS/MS. This approach allowed us to highlight the amino acid sequences and molecular weight distribution of the Ni²⁺-binding peptides, which are critical factors for understanding their metal-binding ability. Besides, using the switch-SENSE® technology, we have acquired unique kinetic data, including affinity constants (k<sub>obs</sub>) and binding dynamics, for both sunflower meal hydrolysates and synthesized model peptides. This innovative applica- tion of switchSENSE® to sunflower meal-derived peptides offers valu- able insights into their interactions with Ni²⁺ ions, significantly advancing the understanding of their binding behaviour and contrib- uting new knowledge to the field. Furthermore, in our study, UV–Vis and CD spectroscopy are used to obtain complementary information on the structural changes and coordination modes involved in Ni²⁺ complex- ation, providing mechanistic insights, currently lacking in the literature. Thus, the combination of these techniques provides a wide range of new information on the structural and biofunctional properties of Ni²⁺- binding peptides from sunflower meal. Our results not only reveal a solid basis for the valorization of this by-product, but also significantly advance the understanding of peptide-metal interactions in food-derived hydrolysates.

## 4. Conclusion and perspectives

Due to the efficiency and selectivity of IMAC technique, SMPI hy- drolysates were enriched in Ni²⁺-binding peptides. All peptide fractions eluted (with imidazole gradient) from the Ni²⁺-IMAC column contained His residue, which could offer a high promising Ni²⁺-binding ability. In all the hydrolysates, several peptide sequences identified using LC-MS/ MS analysis evidenced the presence of His residue at different position from C and N-termini; nevertheless, the most interesting Ni²⁺-binding sequences were characterized by their “ATCUN” motif (i.e., WLH and LLHVT). Indeed, these latter sequences were only found in Prot hydro- lysates (whole and ≤ 1 kDa), suggesting that Prot+Flav sequential treatment could negatively impact the production of potential metal- binding peptides. Moreover, although IMAC technique provided over- all view of Ni²⁺-binding ability of peptides, no statistically significant difference was found between His-containing synthetic peptides by this separation technique. Therefore, thanks to the complementary infor- mation about the Ni²⁺-binding ability of peptides provided by switch-SENSE® technology in static and dynamic modes, we concluded that Pep1.WLHNDGN, 2.WLHNDGNT and 3.FATCIARH showed better Ni²⁺-binding than Pep5.AIPTGTAH, 6.KFPILEH, 7.KFPILEHL and 8. KFPILEHLR, where Pro residue could attenuate such ability. Further- more, the combined application of UV–vis and CD spectroscopic tech- niques allowed to describe the coordination geometry as well as the binding modes of formed peptide-Ni²⁺ complex, which varied according to the nature of the coordinating donor atoms. Overall, the results show that sunflower peptides produced either by hydrolysis or by synthesis,

formed peptide-Ni²⁺ complexes characterized by both octahedral and square-planar geometries. These conformations can be given through nitrogen atoms donor from deprotonated amide and imidazole as well as through the terminal NH₂. Thus, this study provided clear evidence that sunflower meal by-product is a valuable food-derived protein source for the production of Ni²⁺-binding peptides. Peptide sequences highlighted in our study could exhibit a high potential to be used as Cu²⁺-target drug candidates, thus further studies using in vivo system should be carried out to validate their potential.

Finally, note that, the presence of competing ions such as Zn²⁺ in biological or environmental matrices could interfere with Ni²⁺/Cu²⁺ binding and affect the structural integrity of the complexes. Therefore, future studies will aim to address this selectivity factor by evaluating the stability of peptide-metal complexes in the presence of physiologically relevant ions. These investigations would not only improve the under- standing of the stability of the complexes under variable conditions, but also provide valuable insights for their broader practical applications.

### CRediT authorship contribution statement

Jairo Andrés Camaño Echavarría: Writing – original draft, Software, Resources, Methodology, Formal analysis, Data curation, Conceptualization. Christelle Mathé: Writing – review & editing, Methodology. Jean-Michel Girardet: Writing – review & editing, Methodology. Cédric Paris: Writing – review & editing, Methodology, Data curation. Chibuike C. Udenigwe: Writing – review & editing, Methodology. Katalin Selmeczi: Writing – review & editing, Validation, Supervision, Methodology, Data curation. Laetitia Canabady-Rochelle: Writing – review & editing, Visualization, Supervision, Project administration, Funding acquisition, Formal analysis, Conceptualization.

### Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

### Acknowledgments

The authors thank the French ministry government for the MESR grant, the ANR JCJC MELISSA (project number ANR-20-CE43-0002, 2020-2024), the project of the “Lorraine Université d’Excellence” (Investissements d’avenir–ANR project number 15-004). The authors acknowledge technical support in chromatography on the ASIA platform (Université de Lorraine-INRAE) https://a2f.univ-lorraine.fr/en/asia-2/, and the PhotoNS Platform of the L2CM Laboratory, Université de Lorraine.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.jinorgbio.2025.112877.

### Data availability

Data will be made available on request.

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