3.1 Chemical profiles of sugarcane tissues and its fractions
To confirm the oxidative effect on sugarcane plant tissues, some stalk samples were divided and exposed for a long time to visually verify the enzymatic browning. In Fig. 2, it is possible to notice the formation of browned zones in the tissues closest to sugarcane bark. This effect was also noticed in regions closer to the nodes of sugarcane stalk. DESI-MS analyses were carried out on these tissues in order to identify possible chemical compounds that would indicate the identity of the pigments. Although it was not possible to identify the pigment compounds, the analysis by DESI-MS imaging confirmed that there is a large and unaltered distribution of sucrose throughout the transversal surface of the sugarcane stalks. According to Wang et al. (2013), the C-sources like sucrose in sugarcane can be translocated by the distal sink cells or stored in vacuoles throughout the plant cells, i.e., storage parenchyma of sugarcane stalks (Komor, 2000).
Sucrose and its adducts ions are widely found by DESI imaging analysis, such as [sucrose]− (m/z 341), [sucrose + 35Cl]− (m/z 377), [sucrose + 37Cl]− (m /z 379) and [sucrose + HCOO−]− (m/z 387).
Ion m/z 377 (negative mode) and ion m/z 365 (positive mode) fragmentation analyses confirmed the sucrose presence in the sugarcane tissues (Figs. 3 and 4). Fragmentation of the ion m/z 377, negative mode, generated the fragments m/z 179, m/z 161, m/z 113, or even, from the m/z 161, the m/z 131 ion by neutral loss of an aldehyde group (30 Da). The ion m/z 179 has been associated with the formation of monosaccharide moiety (glucose or fructose) from sucrose (m/z 341). Valgimigli et al. (2012) in their studies on the analysis of sugars by HPLC-ESI-MS/MS have demonstrated the formation of fragment m/z 179 referring to the monosaccharide loss, glucose or fructose, leaving another monosaccharide with m/z 179, with subsequent formation of fragments m/z 161 and m/z 113.
Lara-Cruz & Jaramillo-Botero (2022) describe a series of chromatographic and spectrometric analytical methods to identify and quantify sucrose, with LC-MS-ESI being one of these techniques, which we used in this work. Although LC-MS is extensively reported for the secondary compound analyses such as polyphenols, flavonoids, among others, in this work the NMR technique was used to identify the metabolites of the enzymatic browning reaction. As shown in Fig. 5, the analysis of sugarcane tissue presents a diversity of compounds, but due to the high sugar concentrations in its aqueous extracts, the signals referring to phenolic compounds are almost not detected, which required careful analysis of the mass spectra in both mode (negative and positive).
In our previous works, we also noted the difficulty to analyse directly phenolic compounds or flavonoids and its derivatives in sugarcane tissues by ESI-MS (Magri et al., 2019; Sartori et al., 2017; Sartori et al., 2014), because within sugarcane tissues there are high quantities of sugars (15 − 17% wt; Ogando et al., 2022), mainly sucrose (m/z 341) and its monosaccharides (glucose and fructose; m/z 179). These sugars are easily observed by negative mode in ESI-MS. It was shown that signal suppression is most significant for phenolic compounds from sugarcane tissues in the ESI-MS analyses. However, some flavonoids were also identified. The m/z 431 ion was found by ESI-MS analyses and it was identified as apigenin-8-C-glucoside (or only vitexin; Fig. 5), a flavonoid widely found in sugarcane (Colombo et al., 2006; Vila et al., 2008). Other ions were also found and identified as rhamnose moiety (m/z 146), [quinic acid−H-H2O]− (m/z 173), and [guaiacylglyceryl tricin−H]− (m/z 525). However, a molecular ion stood out among the others, the ion for [caftaric acid−H]− (m/z 311). According to Sarni-Manchado et al. (1997), caftaric acid have been associated with oxidation in wines, i.e., high oxidation levels in wines have shown little or no presence of caftaric acid. One important oxidation product derived from caftaric acid has been 2-S-glutathionyl caftaric acid, which can contribute to wine browning. According to Cheynier et al. (1986), the major phenolic compound formed by enzyme oxidation in grapes is 2-S-glutathionyl caftaric acid by mainly proton NMR studies. Condensed tannins (dimers and trimers) have been studied for potential oxidation reactions between, for example, epicatechin-3-o-gallate, allowing the identification of numerous compounds associated with color and aroma in wines (Suc et al., 2021). Yang et al. (2014) reported that the better elucidation of condensed flavonoids was done by LCMS/MS and proton NMR.
Sarni-Manchado et al. (1997) also suggested that m/z 805 (positive mode) and m/z 803 (negative mode) referring to flavylium forms, were associated with the stacking of malvidin-3-O-glucoside in a flavylium form and the caftaric acid (Figs. 5, 6 and 7). Two common forms of favylium cation stacking with caftaric acid have been reported in the literature. We also noticed the presence of m/z 803 ion in the darkened samples of sugarcane tissues (Fig. 6). Figure 7 represents the two most common procyanidin forms formed from stacking of flavylium forms.
Among the widely found sugars attached to flavonoid skeletons, glucose and rhamnose have been widely found (Satterfield & Brodbelt, 2001). Analyzing the aqueous extract samples in positive mode, some flavonoid ions or their condensates were identified, such as: m/z 304 for taxifolin (flavonoid), m/z 595 for cyanidin-3-O-rutinoside and m/z 867 for procyanidin B-type. In a positive mode, the mass spectra were more sensitive to the flavonoid identification than sugars, although there are presence of sodium molecular ions m/z 203 [glucose + Na]+ and m/z 365 [sucrose + Na]+ (Fig. 8).
The widely utilized HPLC-DAD-MS for identifying and quantifying various compounds, including procyanidin B-type trimers (m/z 867) and procyanidin A-type trimers (m/z 865) (Venter et al., 2013). Our analyses indicated the presence of procyanidin B-type trimers (m/z 867) in healthy sugarcane and following enzymatic browning of plant tissues. The presence of cyanidin-3-O-rutinoside molecular ion (m/z 595) and the protonated ion of dihydroxyflavone ([M + H]+; m/z 255) was also noted. The presence of sucrose and glucose (and/or fructose) were also suggested by spectrometry analysis in positive mode; [M + Na]+ (m/z 365 and m/z 203, respectively).
The m/z 430 ion was also found, which has been reported to be α-tocopherol (one carotenoid), but in our studies its identity was not confirmed. Two ions m/z 253 and m/z 284 (negative mode) which present their highest intensities in the oxidized portion of sugarcane tissues, were not identified, unfortunately.
But, no study has been discussed in recent literature on caftaric acid and its derivatives or reaction products in sugarcane, and as an unprecedented work, we used the two analytical techniques to aid us identify the brown compounds formed in the enzyme browning of sugarcane. Since the cost of sugarcane juice treatment due to the juice browning and the impuritie presence has been reported at around R$ 0.23 (Brazilian Real) per ton of sugarcane, i.e., for a sugarcane production equal to 608 million tons, the cost of juice treatment was R$ 139.8 million in 2022 (= US$ 27.9 million).
Thus, colorimetric analyses of total phenolics and flavonoids dosage were carried out to verify their presence and determined their concentrations in sugarcane plant tissues. In order to verify in which sugarcane plant tissues higher concentrations of these phenolic compounds would be found, samples of each sugarcane fraction were analysed separately. The enzymatic activities for the main oxidases found in sugarcane, that is, polyphenol oxidase (or catecholase) and peroxidase as shown in Table 1. The results obtained confirm that the highest concentrations of total phenolics (2,212.8 ± 951.1 mg L− 1) and total flavonoids (53.2 ± 17.6 mg L− 1) are found in the sugarcane top leaf fraction, as well as the highest enzymatic activities for oxidases (PPO: 64.4 ± 2.5 and PDO: 46.3 ± 3.0 U mL− 1). The results showed a statistically significant difference for the Tukey test at p ≤ 0.05. The calibration curves for total phenolics and total flavonoids showed an R2 greater than 0.9000, i.e., R2 = 0.9984 and R2 = 0.9990, respectively. The results for phenolics, flavonoids and enzymatic activities were expressed as the mean of the results of fifteen replicates (3 biological and 5 analytical replicates; n = 15). Once are biological samples collected in agricultural production areas, although randomly, the results showed a higher standard deviation in some analyses. However, as can be seen by the mean comparison analysis (lowercase letters to the right of the results; Table 1) this did not compromise the statistical analysis. On the contrary, given the great disparity in the contents of phenolic compounds and enzymatic activities in each sugarcane fractions, it was possible to confirm that in the top leaves fraction the values were higher than the other fractions, followed by the fraction - nodes.
Table 1
Concentrations of total flavonoids, total phenolics, and polyphenol oxidase and peroxidase enzyme activities from some portions of sugarcane plants.
Samples
|
Total flavonoids
|
Total phenolics
|
Polyphenol oxidase
|
Peroxidase
|
mg L− 1
|
mg L− 1
|
U mL− 1
|
U mL− 1
|
Top leaves
|
53.2±17.6 a
|
2,212.8±951.1 a
|
64.4±2.5 a
|
46.3±3.0 a
|
Nodes
|
5.72±0.61 c
|
70.92±25.7 b
|
34.3±1.8 b
|
28.0±2.2 b
|
Stalks
|
6.87±0.42 b
|
59.51±14.4 b
|
8.7±1.8 c
|
4.7±1.5 c
|
The results were expressed as the mean±standard deviation of 9 replicates: 3 biological and 3 analytical replicates. The lower case letters to right of results represent the significance of the statistical difference between the means for Tukey's 95% test. |
3.2 Reactions of gallic acid by oxidases as model of enzyme browning in sugarcane tissues
Having defined that the sugarcane top leaves contain the highest phenolic compound concentrations and enzymatic activities, in this case, oxidases, we prepared raw enzymatic extracts in potassium phosphate buffer at pH 6.5 to be used as biocatalysts in a model system of enzymatic browning against gallic acid. As the PPO activity was approximately 72% greater than the PDO activity, stability studies and optimal reaction temperature and pH conditions were conducted to determine which would be the optimal temperature and pH for enzymatic browning reactions. From the initial raw enzymatic extract characterization, PPO acitivity and the values found, the results of optimal and stable combinations for the temperature and pH conditions were arrived at, with the combination of the best activity for the enzyme is 30°C at pH 7.0 and the stability combination is around 50°C at pH 5.0 (Fig. 9).
Analyses with oxidases on catechol at three temperatures provided the observation of the increase in absorption over time. Cumulative rates of absorption increasing at 420 nm provided the gradual increasing effect of color in reaction solutions (Fig. 10) and consequent reduction in the gallic acid contents over time, i.e., of 99.0 ± 0 .14 mg L− 1 (control; t = 0) to 48.1 ± 0.14 mg L− 1 (t = 240 min).
To analyse the enzymatic stability of sugarcane oxidases, the synthetic catechol solution was subjected to longer reaction times, initially at 30 min and then 240 min. In both situations, the enzymatic extract of sugarcane oxidases showed to be capable of converting significant amounts of gallic acid into pigment compounds, verified both by the reduction in the concentration of gallic acid by HPLC and by the increase in absorption at 420 nm (Fig. 11). As can be seen, over 30 min of enzymatic reaction the rate of gallic acid conversion was linear at all temperatures tested. The kinetic conversion coefficient of the enzymatic reaction was calculated equal to 0.21 mg L− 1 h− 1 of converted gallic acid.
In conclusion, it was possible to note that in all enzymatic reactions the gallic acid conversion was linear and increasing (Fig. 12), demonstrating that sugarcane oxidases have a great capacity to convert phenolic compounds and there was an intense formation of pigments in reaction systems. From these reaction tests it was not possible to identify which are the possible compounds formed from the gallic acid conversion/degradation by oxidases. Next, NMR and Raman and FTIR spectroscopy analyses were used in an attempt to identify such compounds from the gallic acid conversion.
Polyphenol oxidases are metalloproteins capable of converting polyphenols, generating o-quinones as the main intermediate (Queiroz et al., 2008; Aljawish et al., 2015; Janovitz-Klapp et al., 1990). According to Bolton (2018), o-quinones are highly reactive metabolites derived from catechol in biological matrices when in the presence of oxidases and oxygen. Reactions with amino acids or peptides produce the enzymatic oxidation of phenolic acids such as chlorogenic and caffeic acids (Pierpoint, 1969). Its presence in fruits, vegetables and plants has been associated with an intense brown color (Dulo et al., 2021; Waterhouse & Nikolantonaki, 2015; Ito et al., 2020).
Infrared spectroscopy analysis (Fig. 13) by the percentage of transmittance was possible to identify in region of 1120 cm− 1 an increase in transmittance due to stretching of secondary alcohol C–O group; there was also an increase in 615 cm− 1. However, this variation in transmittance was linked to the stretching of the C–N group and the deformation of the N–H group from amides. There was the disappearance of transmittance in 1450 cm− 1 due to stretching of aromatic group C = C after 60 min of enzymatic reaction. In 1550 cm− 1 was related to appearance of N–H group of acyclic amides (deformation of the N–H bond) after 120 min; between 1120–1100 cm− 1, the FTIR spectrum showed the transmittance to C–OH group of secondary alcohol (stretching of the C–OH bond) after 60 min; 2920 cm− 1, the reason was the appearance of the –CH2– group of alkanes (stretching of the C–H bond), and, finally at 2880 cm− 1, –CHO group of aldehydes appeared as a new band in FTIR spectrum (stretching of the C–H bond).
The analysis by Raman spectroscopy allowed us to identify the chemical alterations that occurred in phenolic acid structures. The structural changes corresponding to action of sugarcane oxidases on gallic acid are sumarized in Table 2. According to data presented in Fig. 14 and Table 2, the decrease in the wavenumbers assigned to the vibrations of C-C aliphatic chains for different exposure times of gallic acid to the enzymatic extract was observed. Additionaly, aliphatic chains of carboxylate salts remained stable in opposition to amide bond vibration wavenumbers, which increased
Table 2
Peak assignment from 1H-NMR spectra of gallic acid solutions against sugarcane enzymatic crude extracts over time.
Group frequencies
|
Wavenumber (cm− 1) / time of enzymatic reaction
|
[Experimental]
|
[Ref.: 28]
|
0
|
60 min
|
120 min
|
180 min
|
240 min
|
C–C aliphatic chains
|
CO2 in plane def vib
|
|
690
|
|
|
|
OH def vib
|
|
837
|
|
|
|
N.D.
|
1047
|
|
|
|
|
Aromatic CH def vib
|
1067
|
1068
|
1063
|
1071
|
1064
|
OH def vib
|
1224
|
1205
|
1212
|
1210
|
1207
|
Carboxylate salts
|
COH bending (phenol)
|
1302
|
1328
|
1339
|
1331
|
1320
|
COH bending (phenol)
|
|
1339
|
|
|
1367
|
N.D.
|
1472
|
|
1464
|
1489
|
|
Amides
|
N.D.
|
|
|
1500
|
|
|
N.D.
|
|
|
1523
|
|
|
C-C stretching
|
|
|
|
|
1620
|
N.D.
|
1634
|
1634
|
|
1638
|
|
N.D.
|
|
|
1645
|
|
1640
|
N.D.
|
1651
|
|
|
1649
|
|
N.D.
|
|
|
|
|
1658
|
C = O stretching
|
|
1666
|
|
|
1683
|
N.D.: not determined; The group frequencies also based on Socrates book (2004) as a suggestion of functional groups and their vibrations. |
According to NMR data, the decreasing at peak intensities with chemical shifts 7.07 ppm and 6.75 ppm, which correspond to hydrogen at positions 2 and 6 of gallic acid and to hydrogen of phenolic groups, respectively, was evidenced in analyses of the NMR data, as illustrated in Fig. 15.
The disappearance of the peaks at 6.24 ppm anda t 7.07 ppm in the region of phenolic acid was observed, after one hour of reaction. We also noted the appearance of the peaks at 7.38 ppm and 2.37 ppm, corresponding to hydrogen from the amine nitrogen and the aliphatic region, respectively. With lower intensity aliphatic expansions of doublets at 2.82 ppm and 2.85 ppm, of the amino acid aspartate, confirm the gallic acid oxidation and subsequent condensation with amino acids, probably side chains or N-terminal from proteins or peptides present in reaction medium, which would reinforce further the formation of pigment compounds. The enzymatic browning of plant tissues, in particular of phenolic acids such as gallic acid, has several reaction stages such as oxidation to quinones and subsequent reaction with amino acids to form highly colored compounds as condensation products (Fig. 16).
The proposed mechanism does not differ from other mechanisms proposed by other authors for other vegetables, such as apples, potatoes, among others. According to Feng et al. (2020) and Liu et al. (2021; 2022), the aspartate amino acid can prevent the discoloration of potato pulp completely and also partly decolorize the brown color, i.e., as we have described, aspartate has been shown to be associated with the browning of sugarcane tissues through an enzymatic browning reaction.