Discovery of a natural cyan blue: A unique food-sourced anthocyanin could replace synthetic brilliant blue

A 3D molecular arrangement and coordination of a minor anthocyanin from red cabbage creates a brilliant blue color.


INTRODUCTION
Despite a long history of exploration, blue remains one of the most challenging colorants to obtain from any source and even more so from natural, edible sources (1)(2)(3)(4)(5)(6)(7). Blue is critical as it is necessary to produce other colors across the palette. Furthermore, the subtleties of color differences in different blues are of great importance in the area of blending colors (8). The two main artificial blue food colorants are brilliant blue (FD&C Blue No. 1) and indigotine (FD&C Blue No. 2), providing cyan ( max = 630 nm) and indigo ( max = 608 nm) hues, respectively. Naturally occurring food-based blues are limited and can be sourced from anthocyanins and a limited set of other blue colorants including phycocyanins (from Spirulina spp.) ( max = 615 to 620 nm) (9) and iridoid derivatives from huito (or gardenia) ( max = 590 to 610 nm) (10,11). However, to date, all known natural colorants have either a  max less than 630 nm, a large violet color contribution (absorbance in the range of 500 to 600 nm), or both (Fig. 1A, method S1.1, and figs. S1.1 and S1.2). Violet contributions affect the final color in blended colorants, e.g., blending natural blues with yellow generally results in a muddy green (fig. S2.1 and discussion S2.1) (8). Although green is abundant in nature, the chlorophyll chromophore is not stable or water soluble (section S2.1) and, therefore, has limited application (12). Achieving a cyan blue from natural sources that could be used as replacement for FD&C Blue No. 1 enabling a broader color palette has been a long-standing challenge to the food industry.
Anthocyanins display a large versatility in color expression because of a complex chemical equilibrium of colored and colorless forms (fig. S3.1 and discussion S3.1) (13)(14)(15)(16)(17). These naturally occurring chromophores are generally intensely red-colored under acidic conditions based on the flavylium cation (Fig. 1B) and change toward violet and blue as the pH increases. Concurrently, the colored forms are susceptible to fading by a combination of water addition (reversible) and autoxidation (irreversible), limiting their stability over time (18). A challenge with anthocyanins is the plethora of different molecules found in a single crop source, such as red cabbage (Fig. 1B) or purple sweet potato (19,20). Both well-established commodity crops generate blue colors at pH 7 to 9. However, it is a composite color because of the numerous individual anthocyanin molecules present and the various equilibrium forms at a given pH for each individual anthocyanin molecule (10,19,20). While the combined red cabbage anthocyanins (RCAs) will produce a vibrant and attractive blue color in pH-neutral solution, there is still a relatively large violet color contribution, limiting their utility as a natural FD&C Blue No. 1 replacement (Fig. 1A).
Previous efforts to identify the chemical structures and spectral properties of RCAs uncovered an interesting structural homology where the common building blocks are a cyanidin chromophore, a sophorose moiety bound at position 3, and a glucose bound at position 5 ( Fig. 1B) (21)(22)(23)(24)(25). The key differentiation factors are the presence of hydroxycinnamic acids and their substitution patterns on sugar 1 (Glc-1) or sugar 2 (Glc-2) of the sophorose (Fig. 1B, table inset). Remarkably, one of the minor mono-acylated anthocyanins, Peak 2 (P2) was found to have a particularly high  max of 640 nm at pH 7 (20). While only a minor component within the RCA mixture, this unique compound warranted further investigation to understand the structural factors conferring its desirable spectral properties.

Structural analysis of P2
Previous efforts relied on mass and retention time or established standards to make predictions of the P2 structure (20). Before conducting an in-depth investigation of the structural factors, which impart the unique spectral properties to P2, it was prudent to obtain a complete structural assignment of the compound. The structural assignment for P2 was determined by high-resolution electrospray ionization-time-of-flight mass spectrometry (ESI-TOF-MS) and a combination of 1 H and 13 C nuclear magnetic resonance (NMR), one-dimensional total correlation spectroscopy (1D TOCSY), correlation spectroscopy (COSY), nuclear Overhauser effect spectroscopy (NOESY), heteronuclear single-quantum correlation spectroscopy (HSQC), and heteronuclear multiple bond correlation (HMBC) experiments and proven to be  (26,27). The results were consistent with the previously predicted structure, supporting that P2 is a regio-isomer of P5 where only the sinapoyl residue resides on a different sugar (Fig. 1B).
It is well established that cyanidin-based chromophores, such as P2, require a combination of neutral pH and complexation to a metal ion, such as an Al 3+ , at the ortho-dihydroxyphenyl moiety (Fig. 1B, positions 3′ and 4′) of the cyanidin chromophore to produce a blue color (15,17,27,28). To evaluate the effect of metal ion complexation with specific RCA components, P2, P5, and P8 were incubated with Al 3+ . The only difference between P2, P5, and P8 is the number and placement of sinapoyl moieties on the sophorose, enabling a direct comparison of the primary structural feature differentiating RCAs and their effect on color formation.
The discrimination between the spectral properties of P2, P5, and P8 in the presence of Al 3+ is highly unexpected ( Fig. 2A). With only one-third equivalent of Al 3+ , the P2 solution gave the desired blue colored complex with a large bathochromic shift of >40 nm ( max at 640 nm; Figs. 1A and 2A, figs. S6.1 to S6.4, and discussion S6.2). In contrast, P8 and P5 had only modest shifts of <20 nm. This provides direct evidence that the sinapoyl moiety and its position (on Glc-2 in P2), despite its intrinsic inability to bind metal ions, play a critical role in the structure of the metal complex, leading to a very strong bathochromic shift.
To elucidate this uniquely colored self-assembled structure of the three P2 molecules coordinated to aluminum metal (29), the circular dichroism (CD) spectrum of P2 and Al 3+ was recorded in buffered solutions. With the addition of Al 3+ , P2 showed an exciton-type positive Cotton effect around  max , while no such Cotton effect was observed without Al 3+ . These results indicate that Al 3+ is critical to the coordination and building of the tertiary structure that exhibits a chiral arrangement of three P2 monomers around the Al 3+ ion (Fig. 3   The UV-visible spectra depicted highlights  max and its respective location on the electromagnetic (EM) absorbance spectra.  max for FD&C Blue No. 1 is 630 nm, spirulina is 617 nm, RCAs at pH 8 are 608 nm, and the Al 3+ (P2 − ) 3 complex at pH 7 is 640 nm. The violet contribution (VC) of each blue, defined as the absorption in the 500-to 600-nm range (depicted in gray), is shown. Values and calculations for VC, i.e., area under the curve (AUC), are provided in sections S1 and S2. (B) The red flavylium cation of the main anthocyanins in red cabbage is depicted as P1 to P8 on the HPLC-diodearray-detection (DAD) trace (520-nm detection) (method S11.2). RCAs display a structural homology (red and blue arrows) where the common building blocks are a cyanidin chromophore (highlighted in red), sugars, and small acyl groups (highlighted in blue). The sophorose moiety is bound at position 3, and a glucose is bound at position 5.
The key differentiation is acyl groups (p-coumaroyl, feruloyl, and sinapoyl) and their substitution patterns on sugar 1 (Glc-1) or sugar 2 (Glc-2) of the sophorose. 1 H NMR measurement of the trimeric complex of Al 3+ (P2 − ) 3 in D 2 O gave a very broad spectrum. However, numbers of signals in the lower field were relatively simple, indicating that the arrangement of the three P2 ligands within the Al complex might be highly symmetrical. Several signals were assignable, but analysis of NOE correlations was difficult (discussion S7.3, figs. S7.11 to S7.15, and table S7.2). Therefore, a structural analysis of Al 3+ (P2 − ) 3 was undertaken by computational modeling. After extensive classical and first-principles molecular dynamics (MD) (9) simulations of P2, plausible geometries were constructed for threefold symmetric propeller-like structures of three P2 molecules around a single aluminum ion and minimized with first-principles calculations (discussion S8.1, method S8.1, fig.  S8.1, and table S8.1). In all proposed configurations, the structure of the P2 cyanidin chromophore is distorted by a torsional displacement , and P8 at pH 7 with and without Al 3+ . The different equivalences of Al are color-coded: black = 0 eq, red = 1 / 3 eq, and blue = 1 eq. The  max values for 0 eq are 598 nm for P2, 596 nm for P5, and 598 nm for P8. Adding 1 / 3 eq Al 3+ generates a  max value of 640 nm for P2, 602 nm for P5, and 617 nm for P8. Adding 1 eq of Al 3+ does not appear to affect  max for P2, P5, or P8. (B) Stability (absorbance at  max of aqueous solutions monitored over time) of anthocyanins P2, P5, and P8 at pH 7 over time with (solid line) and without Al 3+ (dashed line). The addition of Al 3+ enhances the stability of both P2 and P8; however, it has a minor effect on the stability of P5. P8 has an inherent additional stability because of diacylation but is still surpassed by the stability of P2 + Al 3+ . Once the Al 3+ (P2 − ) 3 complex is formed, it demonstrates a marked increase in stability that is thought to stem from the intermolecular interlocking of the P2 moieties (figs. S8.2 to S8.5).  3 were confirmed (details provided in sections S7, S8, and S10). Computational modeling indicates that the large bathochromic shift observed stems mainly from the enhanced distortion of angle , highlighted in blue on the molecular structure. Inset shows the calculated effect of  max as a function of  angle distortion. The color of each circle is the average expected color, the size of the marker corresponds to average oscillator strengths, and error bars are computed as SD of  max . The data for the figure correspond to cyanidin-3-glucoside in fully solvated water. Molecular structures were pruned by replacing the sugar with a methyl group and absorption spectra calculated with time-dependent density functional theory [6-311G(d,p) at B3LYP] (DFT-B3LYP) and fully detailed in section S10. (B) 3D representation and coordination of the anthocyanin P2 around the aluminum.
of the bond between the B and C rings (Fig. 3A,  angle, and fig.  S8.2, inset). A very interesting and remarkable result from the MD simulations, coupled with first-principles evaluation of excited states, is that torsional displacement of this bond causes the greatest change in the simulated color expression (30,31). The - electronic interaction between acyl and cyanidin motifs is not the main cause of the large color shift, as typically suggested. In this case, the HOMO and LUMO energies are more impacted by distortion of the cyanidin nucleus and the interactions among the ones coordinated to the same metal (excitonic effects) than by acyl-cyanidin interactions (discussions S8.1 and S8.2). However, the - interaction between the hydrophobic aromatic rings of the acyl and cyanin groups has a critical influence on the conformations: While intramolecular - stacking dominates in isolated P2 and P5 (discussion S8.3), the proposed Al 3+ (P2 − ) 3 complex manifests intermolecular - stacking between adjacent P2 moieties (interlocking between P2 units shown in figs. S8.2 to S8.4), which is consistent with the strong Cotton effect observed by CD and the enhanced stability of the complex discussed below.

Enzymatic enrichment of P2
While the Al 3+ (P2 − ) 3 complex has the unique chemical conformation that provides the spectral properties desired for replacing FD&C Blue No. 1, P2 represents <5% of the total anthocyanin content naturally occurring in red cabbage. Hence, an ambitious challenge was obtaining sufficient quantities of P2 at high purity for the present research program and for potential use as a food coloring agent. To that end, the homology of structures in red cabbage was leveraged and a hydrolytic enzyme capable of a highly selective catalytic deacylation was found and further developed. The enzyme selectively removes any acyl group bound to Glc-1 of the sophorose moiety while leaving the sinapoyl group on Glc-2 intact, thus converting P6, P7, and P8 into P2 (Fig. 4, A and B). Initial database mining efforts comprised screening a broad range of genes encoding hydrolytic enzymes, including lactonases, esterases, lactamases, and hydrolases, curated from the BRENDA database across 26 enzyme classifications (ECs) (Fig. 4C and tables S11.1 and S11.2) (32). Over three rounds of genomic mining, a total of 46 genes were tested, of which 17 were active on P6 to P8, converting them to P2 (method S11.2). The median sequence identity of the total set of tested enzymes is 7.0%, and the median sequence identity of active enzymes is 13.0% (Fig. 4C), highlighting the plasticity of functionality within the esterase family through the diversity of sequences capable of catalyzing this reaction.
The most active of these enzymes was 1AUR, which in its native form hydrolyzes longer fatty acid chains but has demonstrated broad specificity toward esters ( fig. S11.1) (33). We subsequently redesigned the pocket of the 1AUR protein (24) using a combination of Rosetta Design and FoldIt to introduce new favorable interactions with P8 (the largest and most complex of the RCAs) (34,35). Across multiple rounds of design, the mutant with the best conversion of P6, P7, and P8 was M73H, which was able to drive the transformation of RCAs 3 to 8 to P1 and P2 to completion in a time frame and enzyme concentration viable for the gram-scale production of P2 needed for food-product prototyping (Fig. 4B, table S11.3, and figs. S11.2 and S11.3). On the basis of the modeling and design efforts, we hypothesize that the two additional hydrogen bonds that are developed by the histidine residue substituted at position 73, which are both exposed at the active cleft, strengthen the enzyme-substrate interaction (Fig. 4D).

Color expression and application of P2
Following chromatographic purification of the enzymatic transformation and complex formation with Al 3+ (methods S11.3 to S11.4 and section S12), gram quantities of the Al 3+ (P2 − ) 3 complex were obtained, enabling detailed investigations of the color expression and stability of the novel colorant. In any food product application, color stability is critical. While many naturally sourced colorants, including anthocyanins, have limited stability over time, the Al 3+ (P2 − ) 3 complex at pH 7 showed remarkable stability in sugar syrup over 55 days with only a 14% loss of color (Fig. 5A) (19). Its performance as a colorant was also demonstrated in application to create blue and green colors (hue angle match to a Christmas green shade; fig. S2.1) in several food and confectionery products (Fig. 5, B and C, figs. S13.1 to S13.4, and table S13.1 to S13.4). The stability of this novel colorant in these product applications is excellent as well, with no notable color decay over a 30-day period when stored at ambient conditions (fig. S13.5 and table S13.5). Storage under acidic conditions further enhances the stability of the colorant by reducing fraction of hydrolyzed species, although neutral pH must be able to be restored for integration into product (figs. S3.1 and S9.1) (19). While these initial studies provide a clear starting point for the development of a natural FD&C Blue No. 1, future efforts will be critical to evaluate both stability and color in a wide variety of applications to define usage limitations (discussion S12.1) and appropriate colorant and food safety precautions (36)(37)(38).
In summary, combining modern techniques from analytical chemistry, food science, biochemistry, synthetic biology, color science, and computational chemistry, we found, characterized, and defined a path to production for a naturally sourced cyan blue colorant whose color properties are nearly identical to those of the industry standard brilliant blue FCF (FD&C Blue No. 1). This colorant is also capable of producing superior green colors compared to many existing natural blue colorants. This discovery requiring a global collaboration provides a solution to a long-standing food color need, potentially fulfilling the growing consumer demands for utilization of more natural ingredients in food while keeping a vibrant color palette.

MATERIALS AND METHODS
Detailed methods for the experiments conducted in this study can be found in the Supplementary Materials.

Violet contribution calculation
The violet contribution of a blue colorant was defined as the area under the visible absorption curve (AUC). The area was calculated  Over the course of 10 days, the absorbance at  max of RCAs drops by 57%, indicating loss of cyanidin chromophore, whereas the Al 3+ (P2 − ) 3 complex shows remarkable stability with only 14% loss of absorbance at  max over the course of 55 days. Experimental details are in section S13.3. (B) Sugar-coated lentils (top) using the four blue colorants were obtained under typical panning conditions. The colorants were FD&C Blue No. 1, spirulina, RCAs at pH 8, and Al 3+ (P2 − ) 3 at pH 7. The colorimetry data are plotted in a*b* space and L* values in the sidebar. The Al 3+ (P2 − ) 3 complex at pH 7 provides a very similar hue angle to FD&C Blue No. 1 (sections S1 and S13). (C) Sugarcoated lentils (top) where the colorant blends used were synthetic green, spirulina, red cabbage at pH 8, and Al 3+ (P2 − ) 3 at pH 7, with the latter three mixed with safflower (Saf) as the yellow component. Al 3+ (P2 − ) 3 at pH 7 with safflower provides a very similar hue angle to synthetic green (fig. S13.4 and table S13.4).
using the left Riemann sum for integration, where the height is the absorbance and the width is the spectral resolution (for more specific details, see method S1.1).

Ultraviolet-visible spectrometry
Solutions were poured into relevant cuvette of 10-, 1-, and 0.1-mm cell length, and then ultraviolet (UV)-visible absorption spectra (200 to 800 nm) were recorded on a JASCO V-560 spectrophotometer. The solutions were kept at room temperature and protected from light (method S6.1).

Measurement of CD
CD was measured from 200 to 800 nm with a JASCO J-720 spectrometer using the same solution for the measurement of stability described in section S6.1. To increase signal/noise ratio, the scan was repeated four times, and the data were averaged (method S7.1) (21).

ESI-TOF-MS of aluminum complexes
ESI-MS spectra were recorded with a Bruker COMPACT instrument and analyzed with the application purchased from Bruker Daltonics (method S7.1).

NMR measurement of aluminum complex of P2
NMR spectra were obtained with Bruker Daltonics AVANCE III HD 600 with a TCI cryoprobe and BBO cryoprobe ( 1 H: 600 MHz and 13 C: 150 MHz) in a 5-mm-inside diameter tube at variable temperatures in D 2 O. Chemical shifts were recorded as parts per million (ppm) using the proton resonance in the semi-heavy water (HDO) as a standard (4.67 ppm). Various 1D and 2D measurements were carried out (method S7.3).

Geometric optimization of putative P2 and P5 3:1 complexes with aluminum
Optimizations of either P2 or P5 molecules in a 3:1 complex with aluminum were performed, starting from relevant structures of monomers obtained from extensive MD simulations and analysis, as detailed in section S10.1. These were arranged with a C3 symmetry around Al 3+ and deprotonated at 3′ and 4′, and with Al 3+ coordinated to the oxygens at 3′ and 4′. The two chiral assemblies of the ligands, corresponding to left and right handed (labeled  and , respectively), were tried. Relaxations were then performed in two steps: (i) A preoptimization was run in vacuum with CP2K (39), with DZVP-MOLOPT-SR-GTH basis set and GTH-BLYP pseudopotential (40)(41)(42) corrected with the D3(0) Grimme dispersion (43). (ii) Final optimizations were run in Quantum Espresso (44,45), using ultrasoft pseudopotentials and the Perdew-Burke-Ernzerhof (PBE) exchange-correlation functional (46) with implicit solvent (Environ module) (47) in periodic cubic box of 80 bohr. The following parameters were adopted for the wavefunction convergence: kinetic-energy cutoff of 35 rydberg (Ry) and charge density cutoff of 320 Ry. The Makov-Payne energy correction was used to remove finite-box effects (method S8.1).

Sequence alignment and construction of the phylogenetic tree of esterases of interest
Sequences tested in this study (table S11.1) were aligned using Geneious 2017.10.1.3 using multiple sequence comparison by log-expectation (MUSCLE) alignment (method S11.1.).

Protein purification and activity screening of esterases
An Escherichia coli codon-optimized gene encoding each protein was purchased from Twist Biosciences and transferred into pET29b + to encode a C-terminal hexahistidine tag. Mutant plasmids were produced by Kunkel mutagenesis (55). Plasmids were incorporated into E. coli BL21(DE3) via electroporation. Cultures were grown in Terrific Broth at 37°C, induced with 1 mM isopropyl--d-thiogalactopyranoside, and allowed to express at 18°C for 24 hours, after which cells were lysed, clarified, and protein-purified using gravity columns with immobilized metal affinity chromatography, the details of which are provided in method S11.2. Proteins were screened for activity by combining 10 l of RCE (100 mg/ml) with 90 l of 50 mM Hepes buffer (pH 7.5) and allowed to proceed at room temperature for 24 hours. Reactions were quenched with 70% methanol and brought to pH 3 with 1 l of HCl. Reactions were centrifuged at 4700 rpm for 3 min to remove insoluble protein. Analysis was done using the high-performance liquid chromatography (HPLC) method described in method S11.2.

Gram-scale protein production
The M73H point mutant was created from the 1AUR (sequence ID no. 46) wild-type (WT) plasmid via Kunkel mutagenesis (55) and then transformed into chemically competent E. coli BLR (DE3) cells (details provided in method S11.3). The supernatant lysate containing active enzyme was collected and used for the reaction. The scaled-up generation of P2 via the enzymatic reaction is described in method S11.4.

P2 purification from enzyme-treated red cabbage
The material was purified using several techniques to remove impurities: enzyme precipitation, solid-phase extraction, and preparatory HPLC (more detailed description is found in method S12.1).

Anthocyanin-metal complex [Al 3+ (P2 − ) 3 ] formation details
One-third equivalents of AlK(SO 4 ) 2 stock solution were added to the aqueous P2 solution, adjusted to 7.0. The P2-Al complex solution was transferred to a polypropylene container, immersed in liquid nitrogen until fully frozen, and then placed into the lyophilizer with vacuum set to 0.03 mbar and the condenser to −80°C. More detailed description is provided in method S12.2.