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Zinc-binding behavior of hemp protein hydrolysates: Soluble versus insoluble zinc-peptide complexes
Journal of Functional Foods 49 (2018) 105–112
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Zinc-binding behavior of hemp protein hydrolysates: Soluble versus insoluble zinc-peptide complexes Qingling Wanga, Youling L. Xionga,b, a b
T
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State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu 214122, China Department of Animal and Food Sciences, University of Kentucky, Lexington, KY 40546, United States
A R T I C LE I N FO
A B S T R A C T
Keywords: Hemp protein hydrolysate Peptides Zn2+-binding Bioavailability Isothermal titration
Proteins and peptides when forming complexes with zinc can increase zinc bioavailability. Such complexation was investigated on hemp protein hydrolysates (HPHs) in the present study using Pepsin, Alcalase, Flavourzyme, Papain, Protamex, and Trypsin. Two solubility fractions of Zn2+–HPH complexes, i.e., P1 (water-insoluble large peptides) and P2 (water-soluble small peptides, precipitable by ethanol), were collected. The FTIR analysis on Pepsin-HPH suggested that P1 and P2 peptides had different Zn2+-binding sites where NeH and C]O were the primary sites in P1 and P2, respectively. Although the Zn2+-binding capacity (P1 and P2 combined) of HPHs was lower than that of nonhydrolyzed hemp protein, the P2-bound Zn2+ was more abundant in HPHs (up to 63.4%) than in nonhydrlyzed protein (29.6%). Isothermal titration calorimetry corroborated with Zn2+-binding capacity for different HPH samples. Peptides produced with Flavourzyme had the highest Zn2+-binding activity (88.8%) while those with Pepsin exhibited the maximum solubility.
Chemical compounds studied in this article: Zinc sulfate (PubChem CID: 24424) Silver nitrate (PubChem CID: 24470) Sodium dodecyl sulfate (PubChem CID: 3423265) Acrylamide (PubChem CID: 6579) Ethanol (PubChem CID: 702)
1. Introduction Zinc, the second most abundant inorganic micronutrient found in the human body (Cummings & Kovacic, 2009), is a crucial component within numerous metalloenzymes (Udechukwu, Downey, & Udenigwe, 2018). Moreover, it provides structural integrity to proteins such as transcription factors and hormones (McCall, Huang, & Fierke, 2000). Zinc plays important roles in cell growth and differentiation, protein and DNA synthesis, lipid metabolism, and immune functions (Hambidge, 2000; Ranasinghe et al., 2015). Zinc deficiency can lead to serious health problems, including growth retardation, cognitive impairment, testicular hypofunction, immune dysfunctions, neurological dysfunctions, increased oxidative stress, and increased generation of inflammatory cytokines (Bonaventura, Benedetti, Albarède, & Miossec, 2015; Chasapis, Loutsidou, Spiliopoulou, & Stefanidou, 2012). Zinc is quite mobile from soils and consequently available to plants (Bielicka-Gieldon, Rylko, & Zamojc, 2013). As such, it can be easily incorporated into biogeochemical circulation. Regardless of the source (nuts, seeds, leafy vegetables, animal products, etc.) or form, the bioavailability of zinc depends on its intestinal absorption, which in turn is affected by its release from heterogeneous food matrices (Udechukwu et al., 2018). Some dietary components such as dietary
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fibers, tannins, and phytate can form insoluble complexes with Zn2+, which render the metal unavailable for intestinal absorption (Baye, Guyot, & Mouquet-Rivier, 2017; Kumar, Sinha, Makkar, & Becker, 2010). Zinc supplements such as inorganic zinc salts have been incorporated into food products to prevent zinc deficiency. However, zinc salts can be unstable and cause gastrointestinal tract irritation, making them unsuitable for long-term intake (Akbar, et al., 2013). On the other hand, food-derived peptides can serve as potential Zn2+ carriers in the diet due to their Zn2+-binding capacity (Udechukwu, Collins, & Udenigwe, 2016). Zinc is better absorbed from Zn2+–peptide complexes than from inorganic zinc salts (Udechukwu et al., 2016) or Zn2+–protein complexes (Wang, Zhou, Tong, & Mao, 2011). Zn2+-binding protein hydrolysates and peptides have been obtained from different food protein sources, including milk (Miquel & Farré, 2007; Udechukwu et al., 2018; Wang et al., 2011), chickpea (Torres-Fuentes, Alaiz, & Vioque, 2011), sesame (Wang, Li, & Ao, 2012), oyster (Chen, et al., 2013), silver carp (Jiang, Wang, Li, Wang, & Luo, 2014), rapeseed (Xie et al., 2015), wheat germ (Zhu, Wang, & Guo, 2015), and walnut (Liao et al., 2016). Hemp seed protein has drawn increased commercial attention in recent years due to its purported nutritional value. Hemp protein consists mainly of globulin (edestin) and albumin and has a high digestibility and a well-balanced amino acid composition (Girgih, Udenigwe,
Corresponding author. E-mail address: [email protected] (Y.L. Xiong).
https://doi.org/10.1016/j.jff.2018.08.019 Received 7 April 2018; Received in revised form 1 July 2018; Accepted 8 August 2018 1756-4646/ © 2018 Elsevier Ltd. All rights reserved.
Journal of Functional Foods 49 (2018) 105–112
Q. Wang, Y.L. Xiong
Flavourzyme (pH 7.0, 55 °C), Papain (pH 7.0, 55 °C), Protamex (pH 7.0, 55 °C), Pepsin (pH 2.0, 37 °C), or Trypsin (pH 8.0, 37 °C). The enzyme to HPI substrate ratio was 1:100. After incubation, the enzyme was immediately inactivated by heating in an 85 °C water bath for 10 min. The chilled hydrolysates were brought back to pH 7.0 with 1 N NaOH or 1 N HCl followed by centrifugation at 8000g for 10 min. The supernatants were lyophilized and stored at 4 °C for subsequent analysis.
& Aluko, 2011). The nutritional value of hemp protein has been reported to be comparable to egg white and soybean proteins (Wang, Tang, Yang, & Gao, 2008). Thus, it has great potential to be applied as a valuable food ingredient and nutritional supplementary agent. Numerous studies have indicated that hemp protein is a good source for protein hydrolysates and peptides with substantial ability to scavenge free radicals, chelate metal ions, reduce linoleic acid oxidation, and inhibit ACE activity (Girgih et al., 2011; Orio et al., 2017). However, no study has been conducted on the binding of Zn2+ by hemp protein hydrolysate (HPH) as a possible means to improve the zinc bioavailability as well as expand hemp protein utilization. In most Zn2+–peptide complex studies, peptides in an aqueous solution are incubated with Zn2+ at certain pH, temperature and time conditions; the complexes are subsequently separated through different methods. Udechukwu et al. (2018) and Wang et al. (2011) obtained Zn2+–YCH (yak casein hydrolysate) and Zn2+–WPH (whey protein hydrolysate) complexes through dialyzing the resulting solutions with a semipermeable membrane to remove free Zn2+. Wang, Li, Wang, and Xie (2015) and Xie et al. (2015) recovered the Zn2+–peptides complex by collecting the precipitate after centrifugation at f g. Liao et al. (2016) precipitated Zn2+–walnut peptide complexes in 75% ethanol (1:3, v/v) at 4 °C. Zn2+-binding capacity, calculated based on the zinc content in the complex, may vary depending on the specific separation methods. On the other hand, zinc solubility has a strong correlation with zinc bioavailability. It has been reported that metal-chelating peptides increase mineral bioavailability through maintaining the mineral in a soluble form or increasing its absorption in carrier-mediated processes (Miquel & Farré, 2007).
2.3. Zn2+–HPH binding Zn2+–HPH complexes were prepared and separated according to the method described by Jiang et al. (2014) with some modifications as shown in the following flow chart. HPHs (7.5 mg/mL) were incubated with 9 mM zinc sulfate (ZnSO4) at pH 6.0 and 60 °C for 1 h. After rapidly chilling in an ice slurry, the incubates were centrifuged at 8000g for 15 min to obtain the supernatant (S1) and precipitate (P1). S1 was mixed with ethanol (1:4, v/v), allowed to stand at 4 °C for 1 h, then centrifuged at 8000g for 15 min to yield the supernatant (S2) and precipitate (P2). For comparison, a 4-fold volume of ethanol was added directly to the incubates after reaction followed by centrifugation at 8000g for 15 min to separate the supernatant phase (S3) and precipitate phase (P3). Hence, in terms of protein recovery, these procedures yielded the following results: S1 = S2 + P2; and P3 = P1 + P2. 2.4. Determination of Zn2+-binding capacity To measure the zinc content, samples were first ashed in a Isotemp® Programmable Muffle Furnace (Fisher Scientific, Hampton, NH, USA) at
The objective of the present study was to establish an analytical procedure for the determination of Zn2+-binding capacity of HPHs and to characterize the chemical nature of insoluble versus soluble Zn2+–peptide complexes. Our ultimate goal is to develop technologies for enhancing zinc bioavailability using HPH as the carrier agent.
525 °C for 8 h. The zinc content in P1, P2, P3 and the total zinc content added were measured by an AAnalyst 200 Atomic absorption spectrometer (PerkinElmer Inc., Waltham, MA, USA). The Zn2+-binding capacity was defined as the bound zinc content (in P1 or P2) divided by the total zinc added.
2. Materials and methods
2.5. Fourier transform infrared spectroscopy (FTIR)
2.1. Materials
The stretching or bending of chemical bonds in HPHs due to Zn2+ binding was characterized by a Nicolet Nexus 670 Fourier transform infrared spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The spectra were recorded in the wavenumber region from 600 cm−1 to 4000 cm−1 at a resolution of 4 cm−1 as an average of 64 scans. Significant shifts in absorption bands were used to deduce the possible Zn2+-binding sites.
Hemp protein isolate (HPI) was prepared from raw hemp seeds which were obtained from Yunnan Industrial Hemp Co., Ltd. (Yunnan, China). Hemp seeds were milled and defatted with n-hexane (1:3, w/v) three times. Protein was extracted from the defatted flour dispersed in 15-fold volume (v/w) of water at pH 10.0 and then precipitated by adjusting the pH to 4.5 as described by Tang, Ten, Wang, and Yang (2006). The isoelectric precipitate was suspended in deionized water, adjusted to pH 7.0, then lyophilized. Alcalase, Flavourzyme, and Protamex were purchased from Novozymes North America, Inc. (Franklinton, NC, USA), while Papain, and Trypsin were obtained from Sigma Chemicals (St. Louis, MO, USA). All chemicals of at least a reagent grade were purchased from Sigma Chemicals or Fisher Scientific (Pittsburgh, PA, USA).
2.6. Electrophoresis Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) was performed with a 4% acrylamide stacking gel and a 5–20% acrylamide gradient resolving gel. Protein samples (4 mg/mL) were dissolved in an equal volume of SDS–PAGE sample buffer (4% SDS, 20% glycerol, 0.125 M Tris buffer, pH 6.8). Aliquots of 60 μg of protein per lane were loaded onto the acrylamide gel. After electrophoresis, protein bands were detected through staining either with 1% Coomassie Blue R250 (in 50% methanol and 6.8% acetic acid) or with silver (AgNO3) as described by Swain and Ross (1995).
2.2. Preparation of HPHs HPI (20 mg/mL) was hydrolyzed for 1 h with the following enzymes at their respective optimum conditions: Alcalase (pH 8.5, 55 °C), 106
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from 3270.2 cm−1 to 3289.5 cm−1 (S1) and 3285.8 cm−1 (P1), which may be caused by the strong electron-withdrawing effect of Zn2+ resulting in the repulsion (stretching) of H. The more pronounced NeH shift in S1 than in P1 suggests that water-soluble peptides had a stronger Zn2+-binding ability than less soluble peptides or proteins. Similarly, S2 (the supernatant of ethanol-treated S1) exhibited a stronger NeH stretching than the precipitate (P2), accentuating that soluble peptides were of greater affinity for Zn2+ than less soluble peptides. The absorption band at 1642.7 cm−1 in Pepsin-HPH, characterized as an amide I band, was attributed to the C]O stretching vibration coupled with the bending vibration of NeH (peptide bond) (Chen et al., 2013). This band shifted strongly to 1634.3 cm−1 in P2 and mildly to 1646.2 cm−1 in S1 with no significant change in other samples, indicating that the C]O in P2 was most affinitive of all in coordinating to Zn2+ at this site. The amide II band around 1537.0 cm−1 in Pepsin-HPH, caused by the CeN stretching vibration coupled with NeH bending (Hadnađev et al., 2017), decreased in S1 (1533.8 cm−1) and, not surprisingly, also in its derivative ethanol fractions S2 (1532.8 cm−1) and P1 (1533.0 cm−1). Yet, the shift was not remarkable in P2. On the other hand, the characteristic COO− symmetric stretch at 1396.1 cm−1 moved to greater values in all Zn2+-treated samples, suggesting a strong coordination of Zn2+ and COO−. The peak observed at 1077.8 cm−1, attributed to the stretching of CeO, shifted sharply to 1144.4 cm−1 (S1) with a drastically increased intensity due to the strong effect of the triply degenerated symmetric stretching model of free SO42− (Saha & Podder, 2011). The formation of CeOeZn also contributed to the shift of CeO peak where the wavenumber increased slightly to 1085.4 cm−1 (P1) and 1079.6 cm−1 (P2). It is worth noting that for P3, all its chemical shifts seemed to be the superposition of P1 and P2, as P3 was actually a mixture of P1 and P2. From the above results, it can be inferred that while all the functional peptide groups shown in Fig. 1 were more or less involved in Zn2+ binding, the NeH was the dominant site for the complexation with Zn2+. The results also strongly suggest that soluble peptides played a more important role when compared with insoluble fractions in Zn2+ binding. The oxygen atoms from the carboxyl and carbonyl groups, which were the primary binding sites in P2, were less involved in the binding of Zn2+ by soluble peptides.
2.7. Gel filtration The molecular weights (MWs) of peptides present in S1, S2, and S3 were estimated with a Waters 600 liquid chromatograph system (Waters Co., Milford, MA, USA) equipped with a 2487 UV detector and Empower workstation. A 2000 SWXL TSK gel filtration column (300 mm × 7.8 mm; Tosoh Co., Tokyo, Japan) was used. The mobile phase consisting of acetonitrile/water/trifluoroacetic acid (45/55/0.1, v/v) was set at a flow rate of 0.5 mL/min and the column temperature was kept at 30 °C. Aliquots of 10 µL samples were loaded to the column and the eluents were monitored at 220 nm. A standard MW calibration curve was prepared using cytochrome C (12,500 Da), aprotinin (6,500 Da), bacitracin (1,450 Da), tetrapeptide GGYR (451 Da), and tripeptide GGG (189 Da). 2.8. Isothermal titration calorimetry (ITC) The thermodynamic parameters of titration of ZnSO4 into HPH solutions were determined with a VP-ITC machine (MicroCal, Inc., Northampton, MA, USA) at 60 °C. The machine was carefully calibrated before use. The y-axis was calibrated by sending a series of pulses to the cell heaters, dissipating a known power, and the temperature calibration was done with an internal temperature probe. Both ZnSO4 and HPHs (HPI as the control) were dissolved in 100 mM PIPES buffer (pH 6.1) prior to the titration. The solutions were thoroughly degassed under vacuum prior to measurements to avoid air bubbles in the calorimeter. The number of injections was set to 26 times, and, for each injection, an equal volume (10 µL) of 5 mM ZnSO4 solution was delivered to the sample cell containing 0.5 mg/mL HPHs/HPI over 20 s with an adequate interval (300 s) between injections to allow complete equilibrations. Titrations of ZnSO4 solution into buffer, buffer into HPHs (or HPI) solution, and buffer into buffer under the same conditions were carried out to establish sample blanks. Titrations of buffer into buffer and buffer into HPHs solution caused negligible enthalpy changes. However, the heat change from the instant dispersion of the concentrated ZnSO4 titration drops into buffer could not be ignored. Therefore, experimental data were always corrected by subtracting the data of ZnSO4 solution into buffer to correct for the weak binding of Zn2+ to buffer (Azab, Orabi, & El-Salam, 2001) or buffer dilution. Data were analyzed using Origin 7.0 software supplied by MicroCal. Results were expressed as a plot of enthalpy change per mol of injectant (ΔH, kcal mol−1) versus the ratio of zinc (mmol) to protein (g). The entropy (ΔS) and binding constant (K) were also reported.
3.2. Peptide profiles and molecular weight distribution The protein constituents in HPI and their pepsin hydrolytic products (peptides) in HPH as well as in the supernatants (S1, S2, and S3) and precipitates (P1, P2, and P3) are displayed in Fig. 2. Hemp protein consists of two main fractions, globulin (edestin) and albumin (Wang et al., 2008). Edestin is composed of six identical subunits each made up of an acidic (A) and a basic (B) subunit linked by one disulfide bond (Patel, Cudney, & McPherson, 1994). The ∼34 kDa and ∼21 kDa polypeptides shown in the Coomassie Blue-stained gel were identified as the A and B subunits (Tang et al., 2006; Wang et al., 2008). Pepsin hydrolysis produced peptides with drastically reduced MWs, most of which were less than 21.5 kDa (Fig. 2a). While P1 possessed a relatively broad size distribution (0–45 kDa), the MWs of peptides in S1 were less than 14.4 kDa. Upon the addition of ethanol into S1, most peptides were precipitated (P2), and no clear bands were visible in the supernatant S2 (Fig. 2a). S3, the supernatant collected after precipitating the whole mixture of Zn2+-reacted HPH with ethanol, also showed very faint bands (< 6.5 kDa), consistent with the result of S2. Although Coomassie Blue R-250 is reactive to all proteins, its preference for basic amino acids could potentially lead to an under-detection of peptides. Therefore, silver staining, which is much more sensitive than Coomassie Blue staining, was applied to further visualize peptides in all samples. Although a more intense band (∼6.5 kDa) was seen in S1, still only faint bands were observed in S2 and S3 after silver staining (Fig. 2b). Nevertheless, the A and B subunits of edestin, as well as some large aggregates shown in the upper part of the acrylamide gel in P1, P3, and HPH, were clearly marked by silver, which was in stark contrast to those stained by Coomassie Blue. Yet, very small peptides
2.9. Statistical analysis All experiments were repeated at least three times using different sample preparations, and triplicate analyses were conducted. Analysis of variance (ANOVA) was carried out using Statistix 9.0 (Analytical Software, Tallahassee, FL, USA) in a completely randomized linear model procedure. Significance of differences was defined at P < 0.05 by the least significance difference (LSD) all–pairwise multiple comparisons. 3. Results and discussion 3.1. Characterization of the Zn2+–peptides interaction by FTIR To gain an insight into the functional groups involved in zinc coordination, HPH samples prepared with Pepsin were reacted with ZnSO4, and the resulting infrared spectra of different solubility fractions (supernatants S1, S2, and S3; different precipitates P1, P2, and P3) as well as the non-treated Pespin-HPH control are presented (Fig. 1). The wavenumbers of notable band shifts of these samples are also displayed (see the inset table). Compared with the spectra of Pepsin-HPH, both S1 and P1 obtained after Zn2+ complexation reaction showed significant chemical shifts. In particular, the peak assigned to the stretching vibrations of NeH shifted 107
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100 C–O
Transmittance (%)
90 –OH
80
N–H COO–
70 Pep-HPH S1 S2 P1 P2
60 50 40 4000
3500
C–N C=O N–H N–H
3000
2500
2000
1500
1000
500
-1
Wavenumber (cm )
Sample
N–H stretching
Pep-HPH S1 S2 S3 P1 P2 P3
3270.2 3289.5 3289.9 3287.6 3285.8 3276.4 3279.9
Chemical shifts C=O stretching C–N stretching N–H bending N–H bending 2962.7 1642.8 1537.8 2964.1 1646.2 1533.8 2961.9 1644.0 1532.8 2962.4 1644.9 1533.3 2963.4 1645.7 1533.0 2963.2 1634.3 1535.1 2963.7 1643.1 1536.2
–OH
COO–
C–O
1396.1 1399.0 1398.7 1399.9 1400.5 1402.8 1403.6
1077.8 1144.4 1138.9 1139.3 1085.4 1079.6 1085.9
Fig. 1. FTIR spectra of the supernatants (S1, S2, and S3) and precipitates (P1, P2, and P3) of Pepsin-HPH after incubation with ZnSO4 at 60 °C for 1 h. The table displays the specific chemical shifts where bold-faced values indicate significant changes from Pepsin-HPH (P < 0.05).
Fig. 2. Peptide profile (SDS–PAGE, upper graph) and molecular weight (MW) distribution (gel filtration chromatography, lower table) of the supernatants (S1, S2, and S3) and precipitates (P1, P2, and P3) prepared from Pepsin-HPH after the incubation with ZnSO4 at 60 °C for 1 h. The PAGE was run under non-reducing condition, and protein bands were detected by Coomassie Blue R-250 staining (a) or silver staining (b) where A and B denote, respectively, the acidic and basic subunits of edestin. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
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(a) 200k 116k 97k 66k 45k
shown in Fig. 3a. The enzymatic hydrolysis by all six proteases led to the breakdown of both the acidic (A) and basic (B) subunits of edestin. The bands immediately below 31 kDa in HPI, corresponding to components observed in albumin (Malomo & Aluko, 2015), were also completely hydrolyzed. The ∼48 kDa polypeptide (y), similar to the βsubunit of β-conglycinin in soy protein based on their electrophoretic mobility, was also reported by Tang et al. (2006) and thought to be a 7S-like globulin component in HPI (Wang et al., 2008). This polypeptide was readily degraded by Alcalase, Papain, Protamex, Pepsin, and Trypsin, but somewhat resistant to Flavourzyme. The most prominent band at ∼55 kDa (x) was attributed to the A–B complex (Kim & Lee, 2011; Wang et al., 2008), because its dissociation in the presence of 2mercaptoethanol corresponded to an increased A and B subunit content [see HPI(red)]. The A–B complex was highly susceptible to all enzymes and was ostensibly the source for much of all the low MW peptides shown in the SDS–PAGE pattern. Overall, Pepsin, Papain, and Trypsin were most effective in degrading all HPI components, but Pepsin yielded the most complex mixture of short peptides as also shown previously (Fig. 2). It must be noted that the degradation products would include amino acids and extremely small oligopeptides that were not visible (stainable) in the electrophoretic pattern. The solubility of HPHs corroborated the SDS–PAGE results. Native HPI was not well soluble in water (20.1%), and enzymatic hydrolysis significantly improved (P < 0.05) the solubility by all enzymes (Fig. 3b). The improved solubility was obviously due to the increased number of free (ionizable) amino as well as carboxyl groups derived from extensive peptide bond cleavages or molecular fragmentation, leading to a strong ion–dipole interaction with water. The HPHs prepared with Papain and Pepsin exhibited the highest solubility, 89.9% and 94.2%, respectively, while the HPH with Flavourzyme had the smallest solubility improvement (28.2%). Smaller peptides are expected to have proportionally more polar residues, therefore, an increased propensity to form hydrogen bonds with water to enhance solubility (Gbogouri, Linder, Fanni, & Parmentier, 2004). As a consequence, hydrolysates with smaller peptides were more soluble. The slightly higher solubility of Pepsin-HPH when compared with Papain-HPH may be also related to the fact that the acidic digestion condition for Pepsin (pH 2.0) induced protein structure unfolding (Jiang, Chen, & Xiong, 2009) thereby facilitating the cleavage of susceptible peptide bonds. The ∼48.0 kDa component, reportedly a water-insoluble glycoprotein (Hillestad, Wold, & Engen, 1977) and remained salient in the Flavourzyme-HPH (Fig. 3a), seemed to contribute significantly to the low solubility of the protein digest.
x y
x y A
A
B
B
31k 21.5k 14.4k 6.5k
MW HPI Alc Fla Pap Pro Pep Try HPI(red)
HPHs
(b) 100 Solubility (%)
80
b
c
60
d
40 20
a
a
e f
0 HPI Alc Fla Pap Pro Pep Try HPHs Fig. 3. SDS–PAGE (upper panel) and solubility (lower panel) of hemp protein isolate (HPI) and its hydrolysates (HPHs) prepared with different enzymes: Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try). Samples were not treated with 2-mercaptoethanol except HPI (red) for comparison. A and B denote the acidic and basic subunits of edestin, respectively, and molecular weight (MW) are indicated. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
(< 6.5 kDa), which were detected by Coomassie Blue (Fig. 2a), were not sensitive to silver staining (Fig. 2b). This may be due to different preferences for binding sites between Coomassie Blue and silver. Nielsen and Brown (1984) proposed that silver ions (Ag+) bind only with negatively charged amino acid side chains for color development. These low-MW peptides appeared to be basic as hemp edestin is abundant in arginine (Docimo, Caruso, Ponzoni, Mattana & Galasso, 2014), hence, were more reactive with Coomassie Blue. Gel filtration was applied to qualitatively determine the size distribution of peptide fractions (Fig. 2, inset table). Up to 83.1% peptides of S2 and S3 had a MW lower than 683 Da, namely, they were short peptides comprised of less than five or six amino acid residues. The results were in good correspondence with the faint bands indicated by electrophoresis. Although due to their insolubility, HPH precipitates were not subjected to gel filtration for size analysis, the SDS–PAGE profile suggested that P2 largely consisted of low MW peptides (< 6.5 kDa) (Fig. 2b) when compared with P1.
Fig. 4. Zn2+-binding capacity (combined P1 and P2) and relative zinc content in P2 [i.e., P2/(P1 + P2)] of hemp protein isolate (HPI) and its hydrolysates (HPHs) prepared with different enzymes: Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try). Means (n = 3) of Zn2+ binding rate without a common lowercase letter (a–e), and solubility of complexes without a common uppercase letter (A–D), differ significantly (P < 0.05).
3.3. Comparison of Zn2+ binding of HPHs prepared by different proteases 3.3.1. SDS–PAGE and solubility of HPHs To broaden the investigation of Zn2+–peptide interaction, HPHs were prepared with Alcalase, Flavourzyme, Papain, Protamex, and Trypsin in addition to Pepsin, and the resultant SDS–PAGE profiles are 109
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3.3.2. Zn2+-binding capacity (ZBC) The specificity of enzymes used for proteolysis is an important factor affecting peptide release, while the peptide composition has a great effect on Zn-binding and Zn-solubilization properties of HPHs. The ZBC, which is the combined zinc content in water-insoluble complex P1 (dark grey colored) and in water-soluble complex P2 (light grey colored), i.e., ZBC = P1 + P2, is depicted in Fig. 4. HPI showed the greatest ZBC at 90.7% (with 63.9% and 26.8% attributed to P1 and P2, respectively) when compared with HPHs that ranged from 61.0% to 88.8%. A similar finding was reported by Wang et al. (2011) who claimed that ZBC of native yak casein (92.2%) was significantly higher than that of its hydrolysates (ranging from 32.6 to 59.9%). Drago and Valencia (2004) also noted that zinc dialyzability (release) in a Pepsin digest of casein/Zn mixture increased with casein degradation. It is of interest that of the various protein hydrolysates, the Flavourzyme-HPH, in spite of its low solubility (Fig. 3b), exhibited the highest binding capacity (88.8%) while the Pepsin-HPH, having the greatest solubility (Fig. 3b), had the lowest ZBC (61.0%). Nonetheless, the two HPHs differed sharply in that, P2, which was the ethanol-recovered watersoluble peptide fraction, was largely responsible for the Zn2+ binding in Pepsin-HPH (34.9%), whereas the water-insoluble fraction (P1) was the main contributor to Zn2+ binding in Flavourzyme-HPH (62.8%). The HPHs by other proteases showed more or less equal Zn2+-binding rates between P1 and P2. The relative zinc content in P2 versus P1 is also included (dots) in the figure to provide a comparative view for Zn2+ binding. In spite of the high overall ZBC of HPI, the P2-bound Zn2+ was more abundant in HPHs (up to 63.4%) than in the nonhydrlyzed protein (29.6%). Formation of complexes between peptides and Zn2+ is related to the electron-acceptor potential of Zn2+ and the cation-acceptor ability of peptides as binding ligands, which is pH-dependent. The variation in ZBC between different HPHs may be attributed to sitespecific cleavages that resulted in different peptide profiles (Fig. 3a) and available Zn2+-binding groups, i.e., NeH, C]O, CeN, and COO− (Fig. 1).
Zn2+ to hemp peptides was a dose-dependent endothermic process, which is indicated by the global positive enthalpy change shown in the graph. In particular, the control sample (HPI) produced the strongest endothermic phenomenon through the Zn2+ titration process, followed by Flavourzyme-HPH; Pepsin-HPH had the lowest enthalpy change. The calorimetric data were successfully fitted into the “sequential binding sites” model where the total number of binding sites was set to three in case of overfit (n ≥ 4). The specific thermodynamic parameters, i.e., binding constant (K), enthalpy change (ΔH), and entropy change (ΔS), are tabulated (Table 1). The calculated thermodynamic parameters based on the ITC analysis are conditional parameters, since their values depend on the experimental conditions, such as the pH of the buffer solution and the type of the buffer used (the enthalpy of buffer ionization) (Grossoehme, Spuches, & Wilcox, 2010). The Zn2+ binding to the first and the third sites in the complexation reaction was endothermic because the enthalpy changes (ΔH1 and ΔH3) were both positive. It has been suggested that the endothermic effect due to the dehydration of hydrated Cu2+ could not be overcompensated by the exothermic effect from the formation of new metal–peptide bonds (Wyrzykowski, Zarzeczańska, Jacewicz, & Chmurzyński, 2011). The same thermodynamic subtraction may be applied to the Zn2+–peptide binding in the present study. In addition, the endothermic effect due to the dehydration of peptides (certain polar and charged groups) appeared to also counteract the exothermic metal–peptide bonding effect. Hence, the binding of Zn2+ with hemp peptides on site 1 and site 3 was essentially an entropy driven process, which was confirmed by the ITC (Table 1). A positive value of ΔS is indicative of a significant contribution of non-covalent (ionic) interactions (Makowska et al., 2016). On the other hand, Zn2+ binding to site 2 was found to be exothermic (ΔH < 0), suggesting that this particular locus in hemp peptides had a weak hydration power. Namely, the heat release due to the formation Zn2+–peptide bonds (exothermic) surpassed the absorbed heat when water molecules were liberated from this site. Tang and Skibsted (2016) observed an enthalpy–entropy compensation effect for Zn2+ binding to whey protein, reporting that Zn2+ binding to lactoferrin and bovine serum albumin was exothermic while binding to α-lactalbumin and β-lactoglobulin was slightly endothermic. As a borderline metal ion in the hard/soft classification of Lewis acids, zinc can bind to both nitrogen and oxygen donor ligands as well as sulfur donor ligands. The balance between the softer sulfur ligands and the harder nitrogen ligands in proteins may contribute to the enthalpy–entropy compensation effect. Overall, the calorimetric differences appeared to correspond well with those for ZBC (Fig. 4), indicating that ITC is a valuable method for the characterization and prediction of Zn2+ binding by hemp protein and its enzymatic hydrolysates.
3.3.3. Isothermal titration calorimetry (ITC) To substantiate Zn2+-binding, the HPH samples (and HPI as control) were titrated with ZnSO4 and the calorimetric changes were analyzed by ITC (Fig. 5). The technique has been used to quantitatively estimate the interactions of proteins with metal ions. Overall, the binding of 3.5
HPI Alc Fla Pap HPHs Pro Pep Try
kcal/mole of injectant
3.0 2.5 2.0
4. Conclusions
1.5
Zinc can bind to hydrolyzed hemp protein forming water-soluble (P2) and insoluble (P1) complexes. However, P1 and P2 possessed different Zn2+-binding sites, where NeH groups were the dominant site in P1 and C]O groups were the primary binding site in P2. The P2 fraction mostly consisted of small peptides when compared with P1. Although HPHs had a lower Zn2+-binding capacity than intact hemp protein, they exhibited a greater propensity to form water-soluble Zn2+-peptide complexes, which favor zinc bioavailability. Of the six proteases investigated, Pepsin proved to be most efficient in producing soluble Zn2+–HPH complexes. Understanding the distribution of soluble and insoluble Zn2+–HPH complexes and the physical and thermodynamic nature of the binding, which was achieved through this study, should aid in the application of hydrolyzed hemp protein as a potential ligand for the production of bioavailable zinc.
1.0 0.5 0.0 0.0
0.5
1.0
1.5
2.0
Zn/Protein (mmol/g) Fig. 5. Isothermal titration calorimetry (ITC) of the titration of hemp protein isolate (HPI) and its hydrolysates (HPHs) (0.5 mg/mL) with ZnSO4 (5 mmol/L). A sequential three-site binding model was applied for data fitting. HPHs were prepared with Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try).
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Table 1 Estimated ITC thermodynamic parameters for the interaction of hemp protein isolate (HPI) and its hydrolysates (HPHs) (0.5 mg/mL) with ZnSO4 (5 mmol/L). Zn titration into HPH solution was successfully fitted to a sequential three-site binding model. Data are means ± standard deviations (n = 3).a HPI
K1 ΔH1 ΔS1 K2 ΔH2 ΔS2 K3 ΔH3 ΔS3
8.93 ± 1.60 4.57 ± 0.24 31.8 ± 1.91 0.94 ± 0.35 –22.5 ± 6.50 –53.9 ± 2.59 70.6 ± 22.0 21.4 ± 6.53 86.4 ± 2.57
HPHs Ala
Fla
Pap
Pro
Pep
Try
2.22 ± 0.49 3.29 ± 0.39 25.2 ± 0.43 4.28 ± 1.10 –2.83 ± 0.42 8.12 ± 0.23 7.37 ± 2.10 1.38 ± 0.45 21.8 ± 1.32
4.16 ± 1.20 4.97 ± 0.49 31.5 ± 0.26 1.87 ± 0.80 –11.5 ± 2.89 –19.4 ± 0.54 11.1 ± 5.20 10.1 ± 2.73 48.9 ± 0.35
3.26 ± 0.86 3.06 ± 0.33 25.2 ± 1.34 4.87 ± 1.80 –2.38 ± 0.35 9.73 ± 0.57 3.93 ± 1.80 1.88 ± 0.47 22.1 ± 0.62
1.45 ± 0.43 6.49 ± 1.39 34.0 ± 1.79 15.1 ± 4.60 –5.98 ± 1.36 1.18 ± 0.06 1.86 ± 0.50 2.42 ± 0.39 22.2 ± 0.89
2.67 ± 0.80 1.79 ± 0.26 21.1 ± 0.27 1.07 ± 0.31 –0.94 ± 0.33 11.0 ± 0.25 2.36 ± 0.50 1.10 ± 0.53 18.7 ± 0.98
1.34 ± 0.22 5.85 ± 0.67 31.9 ± 2.20 8.53 ± 1.40 –5.46 ± 0.67 1.60 ± 0.05 0.65 ± 0.10 4.37 ± 0.52 26.0 ± 0.63
HPHs: prepared with Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try). Unit: K (kM−1); ΔH (kcal mol−1); ΔS (cal degree−1). mol a
−1
Acknowledgments
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This study was supported by the USDA National Institute of Food and Agriculture (Hatch project 2351267000) and an Oversea Study Fellowship from the China Scholarship Council (to Q. W.). We thank Dr. Akinbode Adedeji from the Department of Biosystems and Agricultural Engineering at the University of Kentucky for providing the FTIR support. Approved for publication as journal article number 18-07-035 by the Director of the Kentucky Agricultural Experiment Station. Declarations of Interest None. Ethics Statement We declare that the manuscript has been prepared strictly following the high ethics standard of the journal. All the data reported are original and have not been submitted to another journal. References Akbar, B., Niloufar, N., Abolfazl, M., Lofollah, S., Ali, K. Q., & Soheyla, V. (2013). Evaluation and comparison of zinc absorption level from 2-alkyle 3-hydroxy pyranonzinc complexes and zinc sulfate in rat in vivo. Advanced Biomedical Research, 2, 77. Azab, H. A., Orabi, A. S., & El-Salam, E. T. A. (2001). Role of biologically important zwitterionic buffer secondary ligands on the stability of the mixed-ligand complexes of divalent metal ions and adenosine 5’-mono-, 5’-di-, and 5’-triphosphate. Journal of Chemical & Engineering Data, 46, 346–354. Baye, K., Guyot, J.-P., & Mouquet-Rivier, C. (2017). The unresolved role of dietary fibers on mineral absorption. Critical Reviews in Food Science and Nutrition, 57, 949–957. Bielicka-Giełdoń, A., Ryłko, E., & Żamojć, K. (2013). Distribution, bioavailability and fractionation of metallic elements in allotment garden soils using the bcr sequential extraction procedure. Polish Journal of Environmental Studies, 22, 1013–1021. Bonaventura, P., Benedetti, G., Albarède, F., & Miossec, P. (2015). Zinc and its role in immunity and inflammation. Autoimmunity Reviews, 14, 277–285. Chasapis, C. T., Loutsidou, A. C., Spiliopoulou, C. A., & Stefanidou, M. E. (2012). Zinc and human health: An update. Archives of Toxicology, 86, 521–534. Chen, D., Liu, Z., Huang, W., Zhao, Y., Dong, S., & Zeng, M. (2013). Purification and characterisation of a zinc-binding peptide from oyster protein hydrolysate. Journal of Functional Foods, 5, 689–697. Cummings, J. E., & Kovacic, J. P. (2009). The ubiquitous role of zinc in health and disease. Journal of Veterinary Emergency and Critical Care, 19, 215–240. Docimo, T., Caruso, I., Ponzoni, E., Mattana, M., & Galasso, I. (2014). Molecular characterization of edestin gene family in Cannabis sativa L. Plant Physiology and Biochemistry, 84, 142–148. Drago, S. R., & Valencia, M. E. (2004). Influence of components of infant formulas on in vitro iron, zinc, and calcium availability. Journal of Agricultural and Food Chemistry, 52, 3202–3207. Gbogouri, G. A., Linder, M., Fanni, J., & Parmentier, M. (2004). Influence of hydrolysis degree on the functional properties of salmon byproduct hydrolysates. Journal of Food Science, 69, 615–622. Girgih, A. T., Udenigwe, C. C., & Aluko, R. E. (2011). In vitro antioxidant properties of hemp seed (Cannabis sativa L.) protein hydrolysate fractions. Journal of the American Oil Chemists' Society, 88, 381–389. Grossoehme, N. E., Spuches, A. M., & Wilcox, D. E. (2010). Application of isothermal
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Wang, X., Zhou, J., Tong, P., & Mao, X. (2011). Zinc-binding capacity of yak casein hydrolysate and the zinc-releasing characteristics of casein hydrolysate-zinc complexes. Journal of Dairy Science, 94, 2731–2740. Wyrzykowski, D., Zarzeczańska, D., Jacewicz, D., & Chmurzyński, L. (2011). Investigation of copper (II) complexation by glycylglycine using isothermal titration calorimetry. Journal of Thermal Analysis and Calorimetry, 105, 1043–1047. Xie, N., Huang, J., Li, B., Cheng, J., Wang, Z., Yin, J., & Yan, X. (2015). Affinity purification and characterisation of zinc chelating peptides from rapeseed protein hydrolysates: Possible contribution of characteristic amino acid residues. Food Chemistry, 173, 210–217. Zhu, K.-X., Wang, X.-P., & Guo, X.-N. (2015). Isolation and characterization of zincchelating peptides from wheat germ protein hydrolysates. Journal of Functional Foods, 12, 23–32.
Function, 7, 4137–4144. Udechukwu, M. C., Downey, B., & Udenigwe, C. C. (2018). Influence of structural and surface properties of whey-derived peptides on zinc-chelating capacity, and in vitro gastric stability and bioaccessibility of the zinc-peptide complexes. Food Chemistry, 240, 1227–1232. Wang, C., Li, B., & Ao, J. (2012). Separation and identification of zinc-chelating peptides from sesame protein hydrolysate using IMAC-Zn2+ and LC–MS/MS. Food Chemistry, 134, 1231–1238. Wang, C., Li, B., Wang, B., & Xie, N. (2015). Degradation and antioxidant activities of peptides and zinc–peptide complexes during in vitro gastrointestinal digestion. Food Chemistry, 173, 733–740. Wang, X.-S., Tang, C.-H., Yang, X.-Q., & Gao, W.-R. (2008). Characterization, amino acid composition and in vitro digestibility of hemp (Cannabis sativa L.) proteins. Food Chemistry, 107, 11–18.
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Zinc-binding behavior of hemp protein hydrolysates: Soluble versus insoluble zinc-peptide complexes Qingling Wanga, Youling L. Xionga,b, a b
T
⁎
State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu 214122, China Department of Animal and Food Sciences, University of Kentucky, Lexington, KY 40546, United States
A R T I C LE I N FO
A B S T R A C T
Keywords: Hemp protein hydrolysate Peptides Zn2+-binding Bioavailability Isothermal titration
Proteins and peptides when forming complexes with zinc can increase zinc bioavailability. Such complexation was investigated on hemp protein hydrolysates (HPHs) in the present study using Pepsin, Alcalase, Flavourzyme, Papain, Protamex, and Trypsin. Two solubility fractions of Zn2+–HPH complexes, i.e., P1 (water-insoluble large peptides) and P2 (water-soluble small peptides, precipitable by ethanol), were collected. The FTIR analysis on Pepsin-HPH suggested that P1 and P2 peptides had different Zn2+-binding sites where NeH and C]O were the primary sites in P1 and P2, respectively. Although the Zn2+-binding capacity (P1 and P2 combined) of HPHs was lower than that of nonhydrolyzed hemp protein, the P2-bound Zn2+ was more abundant in HPHs (up to 63.4%) than in nonhydrlyzed protein (29.6%). Isothermal titration calorimetry corroborated with Zn2+-binding capacity for different HPH samples. Peptides produced with Flavourzyme had the highest Zn2+-binding activity (88.8%) while those with Pepsin exhibited the maximum solubility.
Chemical compounds studied in this article: Zinc sulfate (PubChem CID: 24424) Silver nitrate (PubChem CID: 24470) Sodium dodecyl sulfate (PubChem CID: 3423265) Acrylamide (PubChem CID: 6579) Ethanol (PubChem CID: 702)
1. Introduction Zinc, the second most abundant inorganic micronutrient found in the human body (Cummings & Kovacic, 2009), is a crucial component within numerous metalloenzymes (Udechukwu, Downey, & Udenigwe, 2018). Moreover, it provides structural integrity to proteins such as transcription factors and hormones (McCall, Huang, & Fierke, 2000). Zinc plays important roles in cell growth and differentiation, protein and DNA synthesis, lipid metabolism, and immune functions (Hambidge, 2000; Ranasinghe et al., 2015). Zinc deficiency can lead to serious health problems, including growth retardation, cognitive impairment, testicular hypofunction, immune dysfunctions, neurological dysfunctions, increased oxidative stress, and increased generation of inflammatory cytokines (Bonaventura, Benedetti, Albarède, & Miossec, 2015; Chasapis, Loutsidou, Spiliopoulou, & Stefanidou, 2012). Zinc is quite mobile from soils and consequently available to plants (Bielicka-Gieldon, Rylko, & Zamojc, 2013). As such, it can be easily incorporated into biogeochemical circulation. Regardless of the source (nuts, seeds, leafy vegetables, animal products, etc.) or form, the bioavailability of zinc depends on its intestinal absorption, which in turn is affected by its release from heterogeneous food matrices (Udechukwu et al., 2018). Some dietary components such as dietary
⁎
fibers, tannins, and phytate can form insoluble complexes with Zn2+, which render the metal unavailable for intestinal absorption (Baye, Guyot, & Mouquet-Rivier, 2017; Kumar, Sinha, Makkar, & Becker, 2010). Zinc supplements such as inorganic zinc salts have been incorporated into food products to prevent zinc deficiency. However, zinc salts can be unstable and cause gastrointestinal tract irritation, making them unsuitable for long-term intake (Akbar, et al., 2013). On the other hand, food-derived peptides can serve as potential Zn2+ carriers in the diet due to their Zn2+-binding capacity (Udechukwu, Collins, & Udenigwe, 2016). Zinc is better absorbed from Zn2+–peptide complexes than from inorganic zinc salts (Udechukwu et al., 2016) or Zn2+–protein complexes (Wang, Zhou, Tong, & Mao, 2011). Zn2+-binding protein hydrolysates and peptides have been obtained from different food protein sources, including milk (Miquel & Farré, 2007; Udechukwu et al., 2018; Wang et al., 2011), chickpea (Torres-Fuentes, Alaiz, & Vioque, 2011), sesame (Wang, Li, & Ao, 2012), oyster (Chen, et al., 2013), silver carp (Jiang, Wang, Li, Wang, & Luo, 2014), rapeseed (Xie et al., 2015), wheat germ (Zhu, Wang, & Guo, 2015), and walnut (Liao et al., 2016). Hemp seed protein has drawn increased commercial attention in recent years due to its purported nutritional value. Hemp protein consists mainly of globulin (edestin) and albumin and has a high digestibility and a well-balanced amino acid composition (Girgih, Udenigwe,
Corresponding author. E-mail address: [email protected] (Y.L. Xiong).
https://doi.org/10.1016/j.jff.2018.08.019 Received 7 April 2018; Received in revised form 1 July 2018; Accepted 8 August 2018 1756-4646/ © 2018 Elsevier Ltd. All rights reserved.
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Flavourzyme (pH 7.0, 55 °C), Papain (pH 7.0, 55 °C), Protamex (pH 7.0, 55 °C), Pepsin (pH 2.0, 37 °C), or Trypsin (pH 8.0, 37 °C). The enzyme to HPI substrate ratio was 1:100. After incubation, the enzyme was immediately inactivated by heating in an 85 °C water bath for 10 min. The chilled hydrolysates were brought back to pH 7.0 with 1 N NaOH or 1 N HCl followed by centrifugation at 8000g for 10 min. The supernatants were lyophilized and stored at 4 °C for subsequent analysis.
& Aluko, 2011). The nutritional value of hemp protein has been reported to be comparable to egg white and soybean proteins (Wang, Tang, Yang, & Gao, 2008). Thus, it has great potential to be applied as a valuable food ingredient and nutritional supplementary agent. Numerous studies have indicated that hemp protein is a good source for protein hydrolysates and peptides with substantial ability to scavenge free radicals, chelate metal ions, reduce linoleic acid oxidation, and inhibit ACE activity (Girgih et al., 2011; Orio et al., 2017). However, no study has been conducted on the binding of Zn2+ by hemp protein hydrolysate (HPH) as a possible means to improve the zinc bioavailability as well as expand hemp protein utilization. In most Zn2+–peptide complex studies, peptides in an aqueous solution are incubated with Zn2+ at certain pH, temperature and time conditions; the complexes are subsequently separated through different methods. Udechukwu et al. (2018) and Wang et al. (2011) obtained Zn2+–YCH (yak casein hydrolysate) and Zn2+–WPH (whey protein hydrolysate) complexes through dialyzing the resulting solutions with a semipermeable membrane to remove free Zn2+. Wang, Li, Wang, and Xie (2015) and Xie et al. (2015) recovered the Zn2+–peptides complex by collecting the precipitate after centrifugation at f g. Liao et al. (2016) precipitated Zn2+–walnut peptide complexes in 75% ethanol (1:3, v/v) at 4 °C. Zn2+-binding capacity, calculated based on the zinc content in the complex, may vary depending on the specific separation methods. On the other hand, zinc solubility has a strong correlation with zinc bioavailability. It has been reported that metal-chelating peptides increase mineral bioavailability through maintaining the mineral in a soluble form or increasing its absorption in carrier-mediated processes (Miquel & Farré, 2007).
2.3. Zn2+–HPH binding Zn2+–HPH complexes were prepared and separated according to the method described by Jiang et al. (2014) with some modifications as shown in the following flow chart. HPHs (7.5 mg/mL) were incubated with 9 mM zinc sulfate (ZnSO4) at pH 6.0 and 60 °C for 1 h. After rapidly chilling in an ice slurry, the incubates were centrifuged at 8000g for 15 min to obtain the supernatant (S1) and precipitate (P1). S1 was mixed with ethanol (1:4, v/v), allowed to stand at 4 °C for 1 h, then centrifuged at 8000g for 15 min to yield the supernatant (S2) and precipitate (P2). For comparison, a 4-fold volume of ethanol was added directly to the incubates after reaction followed by centrifugation at 8000g for 15 min to separate the supernatant phase (S3) and precipitate phase (P3). Hence, in terms of protein recovery, these procedures yielded the following results: S1 = S2 + P2; and P3 = P1 + P2. 2.4. Determination of Zn2+-binding capacity To measure the zinc content, samples were first ashed in a Isotemp® Programmable Muffle Furnace (Fisher Scientific, Hampton, NH, USA) at
The objective of the present study was to establish an analytical procedure for the determination of Zn2+-binding capacity of HPHs and to characterize the chemical nature of insoluble versus soluble Zn2+–peptide complexes. Our ultimate goal is to develop technologies for enhancing zinc bioavailability using HPH as the carrier agent.
525 °C for 8 h. The zinc content in P1, P2, P3 and the total zinc content added were measured by an AAnalyst 200 Atomic absorption spectrometer (PerkinElmer Inc., Waltham, MA, USA). The Zn2+-binding capacity was defined as the bound zinc content (in P1 or P2) divided by the total zinc added.
2. Materials and methods
2.5. Fourier transform infrared spectroscopy (FTIR)
2.1. Materials
The stretching or bending of chemical bonds in HPHs due to Zn2+ binding was characterized by a Nicolet Nexus 670 Fourier transform infrared spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The spectra were recorded in the wavenumber region from 600 cm−1 to 4000 cm−1 at a resolution of 4 cm−1 as an average of 64 scans. Significant shifts in absorption bands were used to deduce the possible Zn2+-binding sites.
Hemp protein isolate (HPI) was prepared from raw hemp seeds which were obtained from Yunnan Industrial Hemp Co., Ltd. (Yunnan, China). Hemp seeds were milled and defatted with n-hexane (1:3, w/v) three times. Protein was extracted from the defatted flour dispersed in 15-fold volume (v/w) of water at pH 10.0 and then precipitated by adjusting the pH to 4.5 as described by Tang, Ten, Wang, and Yang (2006). The isoelectric precipitate was suspended in deionized water, adjusted to pH 7.0, then lyophilized. Alcalase, Flavourzyme, and Protamex were purchased from Novozymes North America, Inc. (Franklinton, NC, USA), while Papain, and Trypsin were obtained from Sigma Chemicals (St. Louis, MO, USA). All chemicals of at least a reagent grade were purchased from Sigma Chemicals or Fisher Scientific (Pittsburgh, PA, USA).
2.6. Electrophoresis Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) was performed with a 4% acrylamide stacking gel and a 5–20% acrylamide gradient resolving gel. Protein samples (4 mg/mL) were dissolved in an equal volume of SDS–PAGE sample buffer (4% SDS, 20% glycerol, 0.125 M Tris buffer, pH 6.8). Aliquots of 60 μg of protein per lane were loaded onto the acrylamide gel. After electrophoresis, protein bands were detected through staining either with 1% Coomassie Blue R250 (in 50% methanol and 6.8% acetic acid) or with silver (AgNO3) as described by Swain and Ross (1995).
2.2. Preparation of HPHs HPI (20 mg/mL) was hydrolyzed for 1 h with the following enzymes at their respective optimum conditions: Alcalase (pH 8.5, 55 °C), 106
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from 3270.2 cm−1 to 3289.5 cm−1 (S1) and 3285.8 cm−1 (P1), which may be caused by the strong electron-withdrawing effect of Zn2+ resulting in the repulsion (stretching) of H. The more pronounced NeH shift in S1 than in P1 suggests that water-soluble peptides had a stronger Zn2+-binding ability than less soluble peptides or proteins. Similarly, S2 (the supernatant of ethanol-treated S1) exhibited a stronger NeH stretching than the precipitate (P2), accentuating that soluble peptides were of greater affinity for Zn2+ than less soluble peptides. The absorption band at 1642.7 cm−1 in Pepsin-HPH, characterized as an amide I band, was attributed to the C]O stretching vibration coupled with the bending vibration of NeH (peptide bond) (Chen et al., 2013). This band shifted strongly to 1634.3 cm−1 in P2 and mildly to 1646.2 cm−1 in S1 with no significant change in other samples, indicating that the C]O in P2 was most affinitive of all in coordinating to Zn2+ at this site. The amide II band around 1537.0 cm−1 in Pepsin-HPH, caused by the CeN stretching vibration coupled with NeH bending (Hadnađev et al., 2017), decreased in S1 (1533.8 cm−1) and, not surprisingly, also in its derivative ethanol fractions S2 (1532.8 cm−1) and P1 (1533.0 cm−1). Yet, the shift was not remarkable in P2. On the other hand, the characteristic COO− symmetric stretch at 1396.1 cm−1 moved to greater values in all Zn2+-treated samples, suggesting a strong coordination of Zn2+ and COO−. The peak observed at 1077.8 cm−1, attributed to the stretching of CeO, shifted sharply to 1144.4 cm−1 (S1) with a drastically increased intensity due to the strong effect of the triply degenerated symmetric stretching model of free SO42− (Saha & Podder, 2011). The formation of CeOeZn also contributed to the shift of CeO peak where the wavenumber increased slightly to 1085.4 cm−1 (P1) and 1079.6 cm−1 (P2). It is worth noting that for P3, all its chemical shifts seemed to be the superposition of P1 and P2, as P3 was actually a mixture of P1 and P2. From the above results, it can be inferred that while all the functional peptide groups shown in Fig. 1 were more or less involved in Zn2+ binding, the NeH was the dominant site for the complexation with Zn2+. The results also strongly suggest that soluble peptides played a more important role when compared with insoluble fractions in Zn2+ binding. The oxygen atoms from the carboxyl and carbonyl groups, which were the primary binding sites in P2, were less involved in the binding of Zn2+ by soluble peptides.
2.7. Gel filtration The molecular weights (MWs) of peptides present in S1, S2, and S3 were estimated with a Waters 600 liquid chromatograph system (Waters Co., Milford, MA, USA) equipped with a 2487 UV detector and Empower workstation. A 2000 SWXL TSK gel filtration column (300 mm × 7.8 mm; Tosoh Co., Tokyo, Japan) was used. The mobile phase consisting of acetonitrile/water/trifluoroacetic acid (45/55/0.1, v/v) was set at a flow rate of 0.5 mL/min and the column temperature was kept at 30 °C. Aliquots of 10 µL samples were loaded to the column and the eluents were monitored at 220 nm. A standard MW calibration curve was prepared using cytochrome C (12,500 Da), aprotinin (6,500 Da), bacitracin (1,450 Da), tetrapeptide GGYR (451 Da), and tripeptide GGG (189 Da). 2.8. Isothermal titration calorimetry (ITC) The thermodynamic parameters of titration of ZnSO4 into HPH solutions were determined with a VP-ITC machine (MicroCal, Inc., Northampton, MA, USA) at 60 °C. The machine was carefully calibrated before use. The y-axis was calibrated by sending a series of pulses to the cell heaters, dissipating a known power, and the temperature calibration was done with an internal temperature probe. Both ZnSO4 and HPHs (HPI as the control) were dissolved in 100 mM PIPES buffer (pH 6.1) prior to the titration. The solutions were thoroughly degassed under vacuum prior to measurements to avoid air bubbles in the calorimeter. The number of injections was set to 26 times, and, for each injection, an equal volume (10 µL) of 5 mM ZnSO4 solution was delivered to the sample cell containing 0.5 mg/mL HPHs/HPI over 20 s with an adequate interval (300 s) between injections to allow complete equilibrations. Titrations of ZnSO4 solution into buffer, buffer into HPHs (or HPI) solution, and buffer into buffer under the same conditions were carried out to establish sample blanks. Titrations of buffer into buffer and buffer into HPHs solution caused negligible enthalpy changes. However, the heat change from the instant dispersion of the concentrated ZnSO4 titration drops into buffer could not be ignored. Therefore, experimental data were always corrected by subtracting the data of ZnSO4 solution into buffer to correct for the weak binding of Zn2+ to buffer (Azab, Orabi, & El-Salam, 2001) or buffer dilution. Data were analyzed using Origin 7.0 software supplied by MicroCal. Results were expressed as a plot of enthalpy change per mol of injectant (ΔH, kcal mol−1) versus the ratio of zinc (mmol) to protein (g). The entropy (ΔS) and binding constant (K) were also reported.
3.2. Peptide profiles and molecular weight distribution The protein constituents in HPI and their pepsin hydrolytic products (peptides) in HPH as well as in the supernatants (S1, S2, and S3) and precipitates (P1, P2, and P3) are displayed in Fig. 2. Hemp protein consists of two main fractions, globulin (edestin) and albumin (Wang et al., 2008). Edestin is composed of six identical subunits each made up of an acidic (A) and a basic (B) subunit linked by one disulfide bond (Patel, Cudney, & McPherson, 1994). The ∼34 kDa and ∼21 kDa polypeptides shown in the Coomassie Blue-stained gel were identified as the A and B subunits (Tang et al., 2006; Wang et al., 2008). Pepsin hydrolysis produced peptides with drastically reduced MWs, most of which were less than 21.5 kDa (Fig. 2a). While P1 possessed a relatively broad size distribution (0–45 kDa), the MWs of peptides in S1 were less than 14.4 kDa. Upon the addition of ethanol into S1, most peptides were precipitated (P2), and no clear bands were visible in the supernatant S2 (Fig. 2a). S3, the supernatant collected after precipitating the whole mixture of Zn2+-reacted HPH with ethanol, also showed very faint bands (< 6.5 kDa), consistent with the result of S2. Although Coomassie Blue R-250 is reactive to all proteins, its preference for basic amino acids could potentially lead to an under-detection of peptides. Therefore, silver staining, which is much more sensitive than Coomassie Blue staining, was applied to further visualize peptides in all samples. Although a more intense band (∼6.5 kDa) was seen in S1, still only faint bands were observed in S2 and S3 after silver staining (Fig. 2b). Nevertheless, the A and B subunits of edestin, as well as some large aggregates shown in the upper part of the acrylamide gel in P1, P3, and HPH, were clearly marked by silver, which was in stark contrast to those stained by Coomassie Blue. Yet, very small peptides
2.9. Statistical analysis All experiments were repeated at least three times using different sample preparations, and triplicate analyses were conducted. Analysis of variance (ANOVA) was carried out using Statistix 9.0 (Analytical Software, Tallahassee, FL, USA) in a completely randomized linear model procedure. Significance of differences was defined at P < 0.05 by the least significance difference (LSD) all–pairwise multiple comparisons. 3. Results and discussion 3.1. Characterization of the Zn2+–peptides interaction by FTIR To gain an insight into the functional groups involved in zinc coordination, HPH samples prepared with Pepsin were reacted with ZnSO4, and the resulting infrared spectra of different solubility fractions (supernatants S1, S2, and S3; different precipitates P1, P2, and P3) as well as the non-treated Pespin-HPH control are presented (Fig. 1). The wavenumbers of notable band shifts of these samples are also displayed (see the inset table). Compared with the spectra of Pepsin-HPH, both S1 and P1 obtained after Zn2+ complexation reaction showed significant chemical shifts. In particular, the peak assigned to the stretching vibrations of NeH shifted 107
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100 C–O
Transmittance (%)
90 –OH
80
N–H COO–
70 Pep-HPH S1 S2 P1 P2
60 50 40 4000
3500
C–N C=O N–H N–H
3000
2500
2000
1500
1000
500
-1
Wavenumber (cm )
Sample
N–H stretching
Pep-HPH S1 S2 S3 P1 P2 P3
3270.2 3289.5 3289.9 3287.6 3285.8 3276.4 3279.9
Chemical shifts C=O stretching C–N stretching N–H bending N–H bending 2962.7 1642.8 1537.8 2964.1 1646.2 1533.8 2961.9 1644.0 1532.8 2962.4 1644.9 1533.3 2963.4 1645.7 1533.0 2963.2 1634.3 1535.1 2963.7 1643.1 1536.2
–OH
COO–
C–O
1396.1 1399.0 1398.7 1399.9 1400.5 1402.8 1403.6
1077.8 1144.4 1138.9 1139.3 1085.4 1079.6 1085.9
Fig. 1. FTIR spectra of the supernatants (S1, S2, and S3) and precipitates (P1, P2, and P3) of Pepsin-HPH after incubation with ZnSO4 at 60 °C for 1 h. The table displays the specific chemical shifts where bold-faced values indicate significant changes from Pepsin-HPH (P < 0.05).
Fig. 2. Peptide profile (SDS–PAGE, upper graph) and molecular weight (MW) distribution (gel filtration chromatography, lower table) of the supernatants (S1, S2, and S3) and precipitates (P1, P2, and P3) prepared from Pepsin-HPH after the incubation with ZnSO4 at 60 °C for 1 h. The PAGE was run under non-reducing condition, and protein bands were detected by Coomassie Blue R-250 staining (a) or silver staining (b) where A and B denote, respectively, the acidic and basic subunits of edestin. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
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(a) 200k 116k 97k 66k 45k
shown in Fig. 3a. The enzymatic hydrolysis by all six proteases led to the breakdown of both the acidic (A) and basic (B) subunits of edestin. The bands immediately below 31 kDa in HPI, corresponding to components observed in albumin (Malomo & Aluko, 2015), were also completely hydrolyzed. The ∼48 kDa polypeptide (y), similar to the βsubunit of β-conglycinin in soy protein based on their electrophoretic mobility, was also reported by Tang et al. (2006) and thought to be a 7S-like globulin component in HPI (Wang et al., 2008). This polypeptide was readily degraded by Alcalase, Papain, Protamex, Pepsin, and Trypsin, but somewhat resistant to Flavourzyme. The most prominent band at ∼55 kDa (x) was attributed to the A–B complex (Kim & Lee, 2011; Wang et al., 2008), because its dissociation in the presence of 2mercaptoethanol corresponded to an increased A and B subunit content [see HPI(red)]. The A–B complex was highly susceptible to all enzymes and was ostensibly the source for much of all the low MW peptides shown in the SDS–PAGE pattern. Overall, Pepsin, Papain, and Trypsin were most effective in degrading all HPI components, but Pepsin yielded the most complex mixture of short peptides as also shown previously (Fig. 2). It must be noted that the degradation products would include amino acids and extremely small oligopeptides that were not visible (stainable) in the electrophoretic pattern. The solubility of HPHs corroborated the SDS–PAGE results. Native HPI was not well soluble in water (20.1%), and enzymatic hydrolysis significantly improved (P < 0.05) the solubility by all enzymes (Fig. 3b). The improved solubility was obviously due to the increased number of free (ionizable) amino as well as carboxyl groups derived from extensive peptide bond cleavages or molecular fragmentation, leading to a strong ion–dipole interaction with water. The HPHs prepared with Papain and Pepsin exhibited the highest solubility, 89.9% and 94.2%, respectively, while the HPH with Flavourzyme had the smallest solubility improvement (28.2%). Smaller peptides are expected to have proportionally more polar residues, therefore, an increased propensity to form hydrogen bonds with water to enhance solubility (Gbogouri, Linder, Fanni, & Parmentier, 2004). As a consequence, hydrolysates with smaller peptides were more soluble. The slightly higher solubility of Pepsin-HPH when compared with Papain-HPH may be also related to the fact that the acidic digestion condition for Pepsin (pH 2.0) induced protein structure unfolding (Jiang, Chen, & Xiong, 2009) thereby facilitating the cleavage of susceptible peptide bonds. The ∼48.0 kDa component, reportedly a water-insoluble glycoprotein (Hillestad, Wold, & Engen, 1977) and remained salient in the Flavourzyme-HPH (Fig. 3a), seemed to contribute significantly to the low solubility of the protein digest.
x y
x y A
A
B
B
31k 21.5k 14.4k 6.5k
MW HPI Alc Fla Pap Pro Pep Try HPI(red)
HPHs
(b) 100 Solubility (%)
80
b
c
60
d
40 20
a
a
e f
0 HPI Alc Fla Pap Pro Pep Try HPHs Fig. 3. SDS–PAGE (upper panel) and solubility (lower panel) of hemp protein isolate (HPI) and its hydrolysates (HPHs) prepared with different enzymes: Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try). Samples were not treated with 2-mercaptoethanol except HPI (red) for comparison. A and B denote the acidic and basic subunits of edestin, respectively, and molecular weight (MW) are indicated. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
(< 6.5 kDa), which were detected by Coomassie Blue (Fig. 2a), were not sensitive to silver staining (Fig. 2b). This may be due to different preferences for binding sites between Coomassie Blue and silver. Nielsen and Brown (1984) proposed that silver ions (Ag+) bind only with negatively charged amino acid side chains for color development. These low-MW peptides appeared to be basic as hemp edestin is abundant in arginine (Docimo, Caruso, Ponzoni, Mattana & Galasso, 2014), hence, were more reactive with Coomassie Blue. Gel filtration was applied to qualitatively determine the size distribution of peptide fractions (Fig. 2, inset table). Up to 83.1% peptides of S2 and S3 had a MW lower than 683 Da, namely, they were short peptides comprised of less than five or six amino acid residues. The results were in good correspondence with the faint bands indicated by electrophoresis. Although due to their insolubility, HPH precipitates were not subjected to gel filtration for size analysis, the SDS–PAGE profile suggested that P2 largely consisted of low MW peptides (< 6.5 kDa) (Fig. 2b) when compared with P1.
Fig. 4. Zn2+-binding capacity (combined P1 and P2) and relative zinc content in P2 [i.e., P2/(P1 + P2)] of hemp protein isolate (HPI) and its hydrolysates (HPHs) prepared with different enzymes: Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try). Means (n = 3) of Zn2+ binding rate without a common lowercase letter (a–e), and solubility of complexes without a common uppercase letter (A–D), differ significantly (P < 0.05).
3.3. Comparison of Zn2+ binding of HPHs prepared by different proteases 3.3.1. SDS–PAGE and solubility of HPHs To broaden the investigation of Zn2+–peptide interaction, HPHs were prepared with Alcalase, Flavourzyme, Papain, Protamex, and Trypsin in addition to Pepsin, and the resultant SDS–PAGE profiles are 109
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3.3.2. Zn2+-binding capacity (ZBC) The specificity of enzymes used for proteolysis is an important factor affecting peptide release, while the peptide composition has a great effect on Zn-binding and Zn-solubilization properties of HPHs. The ZBC, which is the combined zinc content in water-insoluble complex P1 (dark grey colored) and in water-soluble complex P2 (light grey colored), i.e., ZBC = P1 + P2, is depicted in Fig. 4. HPI showed the greatest ZBC at 90.7% (with 63.9% and 26.8% attributed to P1 and P2, respectively) when compared with HPHs that ranged from 61.0% to 88.8%. A similar finding was reported by Wang et al. (2011) who claimed that ZBC of native yak casein (92.2%) was significantly higher than that of its hydrolysates (ranging from 32.6 to 59.9%). Drago and Valencia (2004) also noted that zinc dialyzability (release) in a Pepsin digest of casein/Zn mixture increased with casein degradation. It is of interest that of the various protein hydrolysates, the Flavourzyme-HPH, in spite of its low solubility (Fig. 3b), exhibited the highest binding capacity (88.8%) while the Pepsin-HPH, having the greatest solubility (Fig. 3b), had the lowest ZBC (61.0%). Nonetheless, the two HPHs differed sharply in that, P2, which was the ethanol-recovered watersoluble peptide fraction, was largely responsible for the Zn2+ binding in Pepsin-HPH (34.9%), whereas the water-insoluble fraction (P1) was the main contributor to Zn2+ binding in Flavourzyme-HPH (62.8%). The HPHs by other proteases showed more or less equal Zn2+-binding rates between P1 and P2. The relative zinc content in P2 versus P1 is also included (dots) in the figure to provide a comparative view for Zn2+ binding. In spite of the high overall ZBC of HPI, the P2-bound Zn2+ was more abundant in HPHs (up to 63.4%) than in the nonhydrlyzed protein (29.6%). Formation of complexes between peptides and Zn2+ is related to the electron-acceptor potential of Zn2+ and the cation-acceptor ability of peptides as binding ligands, which is pH-dependent. The variation in ZBC between different HPHs may be attributed to sitespecific cleavages that resulted in different peptide profiles (Fig. 3a) and available Zn2+-binding groups, i.e., NeH, C]O, CeN, and COO− (Fig. 1).
Zn2+ to hemp peptides was a dose-dependent endothermic process, which is indicated by the global positive enthalpy change shown in the graph. In particular, the control sample (HPI) produced the strongest endothermic phenomenon through the Zn2+ titration process, followed by Flavourzyme-HPH; Pepsin-HPH had the lowest enthalpy change. The calorimetric data were successfully fitted into the “sequential binding sites” model where the total number of binding sites was set to three in case of overfit (n ≥ 4). The specific thermodynamic parameters, i.e., binding constant (K), enthalpy change (ΔH), and entropy change (ΔS), are tabulated (Table 1). The calculated thermodynamic parameters based on the ITC analysis are conditional parameters, since their values depend on the experimental conditions, such as the pH of the buffer solution and the type of the buffer used (the enthalpy of buffer ionization) (Grossoehme, Spuches, & Wilcox, 2010). The Zn2+ binding to the first and the third sites in the complexation reaction was endothermic because the enthalpy changes (ΔH1 and ΔH3) were both positive. It has been suggested that the endothermic effect due to the dehydration of hydrated Cu2+ could not be overcompensated by the exothermic effect from the formation of new metal–peptide bonds (Wyrzykowski, Zarzeczańska, Jacewicz, & Chmurzyński, 2011). The same thermodynamic subtraction may be applied to the Zn2+–peptide binding in the present study. In addition, the endothermic effect due to the dehydration of peptides (certain polar and charged groups) appeared to also counteract the exothermic metal–peptide bonding effect. Hence, the binding of Zn2+ with hemp peptides on site 1 and site 3 was essentially an entropy driven process, which was confirmed by the ITC (Table 1). A positive value of ΔS is indicative of a significant contribution of non-covalent (ionic) interactions (Makowska et al., 2016). On the other hand, Zn2+ binding to site 2 was found to be exothermic (ΔH < 0), suggesting that this particular locus in hemp peptides had a weak hydration power. Namely, the heat release due to the formation Zn2+–peptide bonds (exothermic) surpassed the absorbed heat when water molecules were liberated from this site. Tang and Skibsted (2016) observed an enthalpy–entropy compensation effect for Zn2+ binding to whey protein, reporting that Zn2+ binding to lactoferrin and bovine serum albumin was exothermic while binding to α-lactalbumin and β-lactoglobulin was slightly endothermic. As a borderline metal ion in the hard/soft classification of Lewis acids, zinc can bind to both nitrogen and oxygen donor ligands as well as sulfur donor ligands. The balance between the softer sulfur ligands and the harder nitrogen ligands in proteins may contribute to the enthalpy–entropy compensation effect. Overall, the calorimetric differences appeared to correspond well with those for ZBC (Fig. 4), indicating that ITC is a valuable method for the characterization and prediction of Zn2+ binding by hemp protein and its enzymatic hydrolysates.
3.3.3. Isothermal titration calorimetry (ITC) To substantiate Zn2+-binding, the HPH samples (and HPI as control) were titrated with ZnSO4 and the calorimetric changes were analyzed by ITC (Fig. 5). The technique has been used to quantitatively estimate the interactions of proteins with metal ions. Overall, the binding of 3.5
HPI Alc Fla Pap HPHs Pro Pep Try
kcal/mole of injectant
3.0 2.5 2.0
4. Conclusions
1.5
Zinc can bind to hydrolyzed hemp protein forming water-soluble (P2) and insoluble (P1) complexes. However, P1 and P2 possessed different Zn2+-binding sites, where NeH groups were the dominant site in P1 and C]O groups were the primary binding site in P2. The P2 fraction mostly consisted of small peptides when compared with P1. Although HPHs had a lower Zn2+-binding capacity than intact hemp protein, they exhibited a greater propensity to form water-soluble Zn2+-peptide complexes, which favor zinc bioavailability. Of the six proteases investigated, Pepsin proved to be most efficient in producing soluble Zn2+–HPH complexes. Understanding the distribution of soluble and insoluble Zn2+–HPH complexes and the physical and thermodynamic nature of the binding, which was achieved through this study, should aid in the application of hydrolyzed hemp protein as a potential ligand for the production of bioavailable zinc.
1.0 0.5 0.0 0.0
0.5
1.0
1.5
2.0
Zn/Protein (mmol/g) Fig. 5. Isothermal titration calorimetry (ITC) of the titration of hemp protein isolate (HPI) and its hydrolysates (HPHs) (0.5 mg/mL) with ZnSO4 (5 mmol/L). A sequential three-site binding model was applied for data fitting. HPHs were prepared with Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try).
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Table 1 Estimated ITC thermodynamic parameters for the interaction of hemp protein isolate (HPI) and its hydrolysates (HPHs) (0.5 mg/mL) with ZnSO4 (5 mmol/L). Zn titration into HPH solution was successfully fitted to a sequential three-site binding model. Data are means ± standard deviations (n = 3).a HPI
K1 ΔH1 ΔS1 K2 ΔH2 ΔS2 K3 ΔH3 ΔS3
8.93 ± 1.60 4.57 ± 0.24 31.8 ± 1.91 0.94 ± 0.35 –22.5 ± 6.50 –53.9 ± 2.59 70.6 ± 22.0 21.4 ± 6.53 86.4 ± 2.57
HPHs Ala
Fla
Pap
Pro
Pep
Try
2.22 ± 0.49 3.29 ± 0.39 25.2 ± 0.43 4.28 ± 1.10 –2.83 ± 0.42 8.12 ± 0.23 7.37 ± 2.10 1.38 ± 0.45 21.8 ± 1.32
4.16 ± 1.20 4.97 ± 0.49 31.5 ± 0.26 1.87 ± 0.80 –11.5 ± 2.89 –19.4 ± 0.54 11.1 ± 5.20 10.1 ± 2.73 48.9 ± 0.35
3.26 ± 0.86 3.06 ± 0.33 25.2 ± 1.34 4.87 ± 1.80 –2.38 ± 0.35 9.73 ± 0.57 3.93 ± 1.80 1.88 ± 0.47 22.1 ± 0.62
1.45 ± 0.43 6.49 ± 1.39 34.0 ± 1.79 15.1 ± 4.60 –5.98 ± 1.36 1.18 ± 0.06 1.86 ± 0.50 2.42 ± 0.39 22.2 ± 0.89
2.67 ± 0.80 1.79 ± 0.26 21.1 ± 0.27 1.07 ± 0.31 –0.94 ± 0.33 11.0 ± 0.25 2.36 ± 0.50 1.10 ± 0.53 18.7 ± 0.98
1.34 ± 0.22 5.85 ± 0.67 31.9 ± 2.20 8.53 ± 1.40 –5.46 ± 0.67 1.60 ± 0.05 0.65 ± 0.10 4.37 ± 0.52 26.0 ± 0.63
HPHs: prepared with Alcalase (Ala), Flavourzyme (Fla), Papain (Pap), Protamex (Pro), Pepsin (Pep), or Trypsin (Try). Unit: K (kM−1); ΔH (kcal mol−1); ΔS (cal degree−1). mol a
−1
Acknowledgments
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