Hot-Water Infusion as an Efficient and Sustainable Extraction Approach for Edible Flower Teas

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This study investigated the antioxidant effects of tea extracted from flower petals using hot water, suggesting its potential as a natural antioxidant beverage compared to ethanol extracts.

Abstract

This study evaluated hot-water infusion as a practical and sustainable extraction method for functional flower petal teas. Six edible flowers—Tagetes erecta, Lonicera japonica, Celosia argentea var. cristata, Centaurea cyanus, Hibiscus sabdariffa, and Malva sylvestris—were compared under hot-water and 80% ethanol extraction. Hot-water extraction was performed at 100 °C for 15 min. Hot-water extracts showed 1.3–4.0 times higher total phenolic content (TPC) and stronger antioxidant activities (ABTS, DPPH, and FRAP) than 80% ethanol extracts, reflecting efficient extraction of hydrophilic phenolic acids. UPLC-ESI-Q-TOF-MS and GC–MS analyses of hot-water extracts revealed chlorogenic acid, caffeic acid, coumaric acid derivatives, flavonoid glycosides, and aroma volatiles such as hexanal and α-pinene. These findings confirm that simple hot-water infusion effectively recovers both bioactive and aroma-active compounds, supporting its application in developing safe, natural, and functional flower teas.

1. Introduction

Tea is among the most commonly enjoyed drinks across the globe and holds deep cultural and social significance across diverse societies. Beyond its role in daily rituals, tea is valued for its broad spectrum of health benefits, largely attributed to its abundance of bioactive constituents such as polyphenols, flavonoids, and catechins [1,2]. The traditional preparation method—infusing plant materials in hot water—provides a simple yet efficient means of extracting these functional compounds, which confer antioxidant, anti-inflammatory, and other physiological activities. This enduring practice has positioned tea not only as a popular beverage but also as an integral component of a healthy lifestyle.

For many years, edible flowers have been utilized in traditional medicine and culinary applications for their vivid colors, pleasant aromas, and therapeutic properties [3,4]. Building on this traditional use, edible flower teas (EFTs) have recently gained attention as natural, functional beverages that combine aesthetic and flavorful appeal with potential health benefits. As the growing interest among consumers in natural and wellness-oriented products continues to rise, EFTs are emerging as a promising alternative beverage category. Market reports indicate that the global edible flower market is steadily expanding, with a projected CAGR (Compound Annual Growth Rate) of 4.85% from 2021 to 2026 and cumulative growth of 31.17% [4,5]. This growing interest also reflects consumer expectations for safe, clean-label ingredients. To meet these standards, edible flowers are produced under controlled agricultural conditions using organic fertilizers and strict regulations on pesticide residues, and are grown in soils free of heavy metals and other contaminants [4].

Like conventional teas, flower petals contain diverse phytochemicals, including phenolic compounds and volatile organic compounds (VOCs), that contribute to their color, aroma, flavor, and biological activities. Conventional herbal tea contains polyphenols dominated by catechins and theaflavins, whereas edible flower teas are characterized by high levels of flavonol glycosides such as quercetin, kaempferol derivatives, and rutin [2,6]. Although the phytochemical profiles of herbal teas and flower teas differ, both are well known for their beneficial bioactivities, including antioxidant and anti-inflammatory effects. Phenolic compounds, such as phenolic acids, flavonoids, and their glycosidic derivatives [3], are a major class of plant secondary metabolites characterized by one or more hydroxyl groups attached to aromatic rings. Phenolics play a critical role in human health due to their strong antioxidant capacity, which mitigates oxidative stress and inflammation, key factors associated with chronic diseases such as cancer, diabetes, and cardiovascular disorders [7]. In addition to phenolics, VOCs, including terpenoids, phenylpropanoids, and fatty acid derivatives, impart characteristic floral aromas and exhibit antimicrobial and antioxidant properties [8]. These bioactive constituents collectively contribute to the dual functionality of EFTs as health-promoting ingredients and natural flavoring agents in functional foods and beverages. Recent studies have further clarified that these bioactive constituents provide meaningful health benefits, particularly antioxidant and anti-inflammatory effects [6,9]. Such findings support the classification of edible flower teas as functional foods and highlight the importance of characterizing their extraction behavior and composition under consumption-relevant conditions.

However, a key question remains as to whether traditional hot-water infusion effectively extracts these bioactive components from flower petals. Although organic solvents such as ethanol are typically used in analytical and pharmaceutical fields to maximize the yield of functional compounds [1,5,10], this efficiency advantage comes with notable environmental concerns, including solvent toxicity, disposal issues, and energy-intensive processing [11]. To address these limitations, various green extraction technologies such as subcritical water extraction, ultrasound-assisted extraction, pressurized liquid extraction, pulsed electric fields, microwave-assisted extraction, and supercritical fluid extraction have been introduced as more sustainable alternatives to conventional organic solvent methods [12,13]. Nevertheless, these technologies do not reflect the real-world consumption context in which edible flowers are typically prepared and consumed as simple hot-water infusions. Thus, the novelty of this study lies in directly comparing conventional aqueous infusion with standard ethanolic extraction to validate the practicality and functional potential of EFTs.

Therefore, conventional hot-water infusion of six edible flower petals—Tagetes erecta (Marigold), Lonicera japonica (Honeysuckle), Celosia argentea var. cristata (Cockscomb), Centaurea cyanus (Cornflower), Hibiscus sabdariffa (Roselle), and Malva sylvestris (Common Mallow)—was compared with ethanolic extracts coupled with homogenization and ultrasonication, which are generally used as food-grade organic solvent extraction methods, for evaluating total phenolic content and antioxidant capacities. Furthermore, to elucidate the compositional characteristics of EFTs, phytochemical profiling of the hot-water extracts was analyzed using liquid chromatography coupled with tandem mass spectrometry (LC–MS/MS), and volatile constituents were characterized by gas chromatography–tandem mass spectrometry (GC–MS/MS). Through this integrated analytical approach, this study aims to demonstrate that traditional hot-water infusion can effectively extract key bioactive compounds, thereby establishing EFTs as a scientifically supported functional beverage.

2. Materials and Methods

2.1. Chemicals and Materials

The dried edible flowers of six species (Tagetes erecta (TE), Lonicera japonica (LJ), Celosia argentea var. cristata (CA), Centaurea cyanus (CC), Hibiscus sabdariffa (HS), and Malva sylvestris (MS)) were purchased from an online market in the Republic of Korea. The identity of each flower species was confirmed based on its morphological characteristics (color, petal shape, and size), which were compared with descriptions provided in previous studies [14,15,16,17,18,19]. Samples were placed at room temperature (Figure 1). The chemical solvents (ethanol and hydrochloric acid) were purchased from Daejung (Siheung-si, Republic of Korea). The reagents included Folin & Ciocalteu’s phenol, DPPH (2,2-Diphenyl-1-picrylhydrazyl), TPTZ (2,4,6-tri [2-pytidyl]-s-triazine), AAPH (2,2′-Azobis-(2-AmidinoPropane) dihydrochloride), ABTS (2,2′-Azino-Bis(3-ethylbenzThiazoline-6-Sulfonic acid)diammonium), FeCl₃, FeSO₄·7H₂O, gallic acid, and ascorbic acid, which were purchased from Sigma Aldrich (St. Louis, MO, USA).

2.2. Extraction of Bioactive Compounds from EFTs by Hot-Water Infusion and Ethanol Extraction

2.2.1. Hot-Water Extraction

Hot-water extraction was adopted to reflect actual consumption. Two hundred milliliters of deionized water (DIW) was heated to 100 °C using a hot plate with a magnetic stirrer. Once the desired temperature was reached, 1 g of the sample (Figure 1) was added to the heated DIW and extracted for 15 min. Each sample was filtered using No. 1 Whatman filter paper, and the final extract volume was brought to 200 mL using DIW. The samples were acidified with 0.1% hydrochloric acid (HCl) to stabilize pigments [20]. The extracts were stored at −20 °C for further studies.

2.2.2. Ethanol Extraction

Ethanol extraction was used as a comparative method to hot-water extraction, following a widely accepted standard based on a previous reference [21]. After grinding six species of flower samples (Figure 1) using a blender (SBL316ABD, Home Planet, Seoul-si, Republic of Korea), each EFT sample (3.5 g) was mixed with 140 mL of 80% aqueous ethanol. The mixture was homogenized at 12,000 rpm for 2 min. Subsequently, the mixture was subjected to ultrasonic extraction using an ultrasonic cleaner set (Digi-10H, SciLab, Seoul-si, Republic of Korea) for 30 min. The extract was filtered using a vacuum filtration with No. 1 Whatman filter paper. This extraction process was replicated twice. Following ethanol extraction, a total volume of 420 mL of extract was obtained. The extracts were added to 0.1% HCl to preserve the phenolics and stored at −20 °C.

2.3. Comparative Evaluation of Total Phenolic and Flavonoid Contents in Hot-Water Infusion and Ethanol Extracts

2.3.1. Total Phenolic Content (TPC)

The total phenolic content assay was conducted using a spectrometer according to the method of Baek et al. [22]. A total of 10 μL of each properly diluted sample was mixed with DIW (130 μL). Subsequently, 10 μL of Folin & Ciocalteu’s phenol reagent was added into the reaction mixture. After 6 min at room temperature for reaction, 100 μL of 7% Na₂CO₃ solution was dispensed into each well. Upon completion of the 90 min reaction, the absorbance was monitored at 750 nm using the Cytation 5 cell imaging multimode reader (Agilent Biotek, Santa Clara, CA, USA). The standard of gallic acid solution was diluted to 10, 30, 60, and 100 μg/mL, and the TPC was reported as mg of gallic acid equivalent (GAE) per g of dry matter. The regression equation (y = 0.0031x + 0.0005) was obtained, and the correlation coefficient (R² > 0.99) confirmed excellent linearity across the tested concentration ranges.

2.3.2. Total Flavonoid Content (TFC)

The total flavonoid content was analyzed via spectrometric measurement at 430 nm. In total, 50 μL of each EFT extract was mixed with 100 μL of methanol along with 20 μL of 10% aluminum chloride solution in a 96-well microplate. After 3 min, 20 μL of sodium acetate and 60 μL of methanol were introduced. The reaction mixture was reacted in a dark room for 40 min at room temperature. The standard curve of quercetin was diluted to 10, 20, 30, and 40 μg/mL, and results were expressed as milligrams of quercetin equivalent (QE) per gram of dry weight. A calibration curve described by the equation y = 0.0093x − 0.0041 was generated, and the strong correlation (R² > 0.99) demonstrated outstanding linearity within the evaluated concentration range.

2.4. Determination Antioxidant Activity

2.4.1. DPPH Radical Scavenging Activity Assay

Here, 0.1 mM of DPPH solution in 80% methanol was prepared and left in a dark room for 20 min. The absorbance of the methanolic DPPH radical solution was adjusted to 0.700 ± 0.010 at 517 nm using 80% methanol. Then, 5 μL of each sample and aliquots of the standard solution were dispensed into a 96-well plate, after which 245 μL of the methanolic DPPH solution was subsequently added. The mixture was reacted in the dark room for 30 min, and the absorbances were measured at 517 nm. Ascorbic acid served as the reference compound, with standard solution prepared at 10, 30, 60, and 100 μg/mL. Results were expressed as milligrams of vitamin C equivalent (VCE) per gram of dry weight. The calibration data yielded the regression model y = 0.0038x − 0.0009, with an R² value exceeding 0.99, indicating highly linear behavior over the tested concentration interval.

2.4.2. ABTS Radical Scavenging Activity Assay

The ABTS working solution was generated by reacting 1.0 mM AAPH with 2.5 mM ABTS in an amber container filled with 50 mL of PBS. The reaction mixture was incubated in an 80 °C water bath for 40 min, followed by cooling to ambient temperature. After filtration through a 0.45 μm PVDF membrane, the solution was adjusted with PBS until its absorbance reached 0.700 ± 0.010 at 734 nm. Next, 10 μL of each standard and samples were mixed with 240 μL of the diluted ABTS reagent and left at 37 °C for 10 min. Absorbance readings were recorded at 734 nm. Ascorbic acid standards were prepared in a dilute HCl aqueous medium, expressed as mg VCE/g dw. From the standard curve, the regression equation y = 0.0021x + 0.0037 was obtained, and the correlation coefficient (R² > 0.99) verified excellent linearity throughout the measurement range.

2.4.3. Ferric Reducing Antioxidant Power (FRAP) Assay

The FRAP assay was performed according to a modified method using 300 mM acetate buffer (pH 3.6), 10 mM TPTZ (in 40 mM HCl), and FeCl₃·6H₂O (20 mM) mixed at a 10:1:1 (v/v/v) ratio. A 5 μL aliquot of sample extract was introduced into 250 μL of pre-warmed FRAP reagent (37 °C). Absorbance measurements were obtained at 593 nm after 4 min. The standard solution was prepared by diluting FeSO₄·7H₂O solution to concentrations of 0–800 μM, and the results were shown as μM FeSO₄ equivalent/g dw. The resulting standard curve followed the regression equation y = 0.2906x − 0.0049, and the high determination coefficient (R² > 0.99) confirmed robust linearity across the tested concentration levels.

2.5. Identification of Phenolic Acids and Flavonoids by UPLC-ESI-Q-TOF-MS

Hot-water extracts were examined using an UPLC system coupled to an ESI-enabled quadrupole time-of-flight mass spectrometer (SYNAPT XS, Waters, Milford, MA, USA). Separation was performed on an ACQUITY UPLC BEH C18 column (Waters, USA; 100 × 2.1 mm, 1.7 μm) held at 40 °C. Dual-mode (ESI⁺/ESI⁻) comparative metabolomic analysis was used to establish complementary profiles of phenolic acids and flavonoids of the samples. In positive ionization mode, 1 μL of the sample was injected, while in negative ionization mode, 2 μL was injected. The mobile phase was delivered at a flow rate of 0.3 mL/min. The mobile phase consisted of (A) deionized water containing 0.1% formic acid and (B) acetonitrile containing 0.1% formic acid. The gradient conditions were established as follows: 0–1 min, 0% B; 1–25 min, a linear increase to 100% B; 25–27 min, 100% B; 27–27.01 min, return to 0% B; and 27.01–30 min, equilibration at 0% B. The total running time was 30 min. The ESI capillary and cone voltages were set at 2 kV and 40 V, respectively. The source and desolvation temperatures were maintained at 100 °C and 250 °C, respectively. The cone gas flow rate was 50 L/h, desolvation gas was 600 L/h, and nebulizer pressure was 6.5 bar. Full mass spectra were collected in both ionization modes over an m/z range spanning 100 to 1100 Da. Raw spectral data obtained from both positive and negative ionization modes were processed, normalized, and retention time aligned using Progenesis QI software 2.4 (Waters, Milford, MA, USA) for dual-mode comparative metabolomic analysis. Metabolite annotation was performed based on accurate mass measurement, isotopic distribution, and fragmentation pattern comparison with public databases (chemspider) and previously reported edible flower compounds. No authentic standards were used in this study.

2.6. Identification of Volatile Organic Compounds by HS-SPME/GC-MS

Volatile organic compounds were analyzed using headspace solid-phase microextraction using an autosampler coupled with gas chromatography–mass spectrometry (HS-SPME/GC–MS) (Nexis GC-2030, MS-QP2020NX, and HS-20 NX, Shimadzu, Kyoto, Japan). Due to the sensory differences in the aroma of flower teas by the type and condition of samples, extraction process, and brewing conditions, in this study, HS-SPME analysis was performed on the dried plant materials under hot-water preparation-like conditions rather than on the final aqueous extracts. Therefore, the identified VOCs should be interpreted as potential aroma components inherent to the dried flower materials that are releasable during hot-water preparation rather than as volatiles directly quantified in the brewed infusions. Here, 1 g of each sample was placed in a 20 mL vial and heated in a dry oven at 100 °C for 15 min to allow for preconditioning. The headspace extraction was performed in loop mode at 80 °C with an equilibration time of 15 min. Chromatographic separation was carried out using a DB-1 fused-silica capillary column (60 m × 0.32 mm i.d., 1.00 μm film thickness; J&W DB-1, Agilent Technologies, USA), with helium as the carrier gas. The injection volume was 1 mL in split mode. The GC oven was programmed starting at 35 °C with a 10 min hold, ramped to 120 °C at 8 °C/min and held for another 10 min, then raised to 180 °C at 12 °C/min with a 7 min hold, and finally increased to 230 °C at 15 °C/min with a concluding 5 min hold. The total running time was 50.96 min.

Analyses were conducted using electron-impact (EI) ionization at 70 eV, with a scan rate of 20,000 amu/s. The performance of the mass spectrometer was verified using 1 pg of octafluoronaphthalene (OFN, m/z 272), ensuring a signal-to-noise ratio (S/N) ≥ 2000. Compound identification was conducted by comparing the acquired spectra with the NIST (Wiley, Hoboken, NJ, 107, and 147) mass spectral libraries. The ion source temperature was maintained at 220 °C. Raw data were processed using GC-MS Post-run Analysis software 4.45 (Shimadzu, Kyoto, Japan).

2.7. Statistical Analysis

All experiments were performed with three independent replicates. One-way ANOVA and t-tests were carried out using GraphPad Prism 10 (Graphpad Software, SanDieogo, CA, USA). Differences were regarded as statistically meaningful when the p-value was below 0.05. Also, heatmap visualization was performed using GraphPad Prism 10. Analyses were conducted using raw data without normalization or transformation, allowing for direct comparison of the relative trends among measured parameters.

3. Results

3.1. Comparison of Extraction Efficiency Between Hot-Water Infusion and 80% Ethanol Extracts

Table 1. Comparison ¹ of total phenolic contents (TPCs) and total flavonoid contents (TFCs) between hot-water extracts and 80% ethanol extracts.

Sample Code 2	TPC (mg GAE/g dw)		TFC (mg QE/g dw)
	Hot-Water	80% Ethanol	Hot-Water	80% Ethanol
TE	64.72 ± 0.71 a***	49.66 ± 4.60 a	21.37 ± 1.46 a***	25.72 ± 1.06 a
LJ	59.13 ± 1.02 b***	40.37 ± 0.58 b	3.51 ± 0.85 b**	2.28 ± 0.27 c
CA	18.84 ± 0.92 cd***	8.07 ± 0.22 c	2.14 ± 0.16 c***	4.19 ± 0.15 b
CC	16.05 ± 1.24 d***	3.98 ± 0.11 e	2.57 ± 0.42 bc***	0.96 ± 0.04 e
HS	17.28 ± 5.41 d***	7.95 ± 0.16 c	2.42 ± 0.52 c*	1.80 ± 0.11 cd
MS	22.12 ± 2.09 c***	6.69 ± 0.19 d	1.54 ± 0.19 c	1.54 ± 0.06 de

¹ Different lowercase letters indicate significant differences within a column among TE, LJ, CA, CC, HS, and MS, as determined by one-way ANOVA (p < 0.05). Statistical significance within a row between hot water and 80% ethanol is indicated as *** (p < 0.0001), ** (p < 0.001), and * (p < 0.05) according to the t-test. ² Details of the sample code are provided in Figure 1.

summarizes the TPC and TFC of EFTs extracted using hot water and 80% ethanol. In terms of total phenolic content (TPC), the hot-water extracts exhibited markedly higher extraction efficiency than the 80% ethanol extracts, whereas the recovery of flavonoids (TFC) showed only minor differences between the two solvents. Among the hot-water extracts, TE contained the highest TPC (64.72 ± 0.71 mg gallic acid equivalent/g dw), whereas CC exhibited the lowest (16.05 ± 1.24 mg GAE/g dw). The 80% ethanol extracts also followed a similar pattern but with generally lower values than hot-water extracts, with TE showing the highest TPC (49.66 ± 4.60 mg GAE/g dw) and CC the lowest (3.98 ± 0.11 mg GAE/g dw). These results indicate that hot water was more effective at releasing phenolic compounds, likely due to enhanced solubilization of hydrophilic phenolics under elevated temperatures. Although the 80% ethanol extracts yielded slightly higher TFC values in a few samples (e.g., TE and CA), the overall magnitude of TFC between the two solvents remained relatively comparable. The highest TFC among hot-water extracts was also observed in TE (21.37 ± 1.46 mg quercetin equivalent/g dw), while MS exhibited the lowest (1.54 ± 0.19 mg QE/g dw). In 80% ethanol extracts, TE and CC again showed the highest (25.72 ± 1.06 mg QE/g dw) and lowest (0.96 ± 0.04 mg QE/g dw) values, respectively. Overall, these data suggest that hot-water infusion provides more efficient recovery of polyphenolic compounds, which may contribute to enhanced antioxidant activities.

3.2. Comparison of Antioxidant Activity Between Hot-Water Extracts and 80% Ethanol Extracts

Table 2. Comparison ¹ of ABTS radical scavenging activity, DPPH radical scavenging activity assay, and Ferric reducing antioxidant power (FRAP) assay between hot-water extracts and 80% ethanol extracts.

Sample Code 2	ABTS (mg VCE/g dw)		DPPH (mg VCE/g dw)		FRAP (μM FeSO4/g dw)
	Hot-Water	80% Ethanol	Hot-Water	80% Ethanol	Hot-Water	80% Ethanol
TE	101.41 ± 4.15 a*	106.23 ± 3.48 a	62.44 ± 4.73 b*	57.08 ± 5.93 a	1345.91 ± 118.43 a*	1251.94 ± 54.01 a
LJ	45.98 ± 2.13 b*	49.11 ± 2.81 b	66.56 ± 4.71 a***	22.80 ± 5.39 b	812.91 ± 63.68 b***	553.17 ± 10.66 b
CA	34.32 ± 5.30 c***	9.98 ± 0.43 c	9.48 ± 0.42 c	9.75 ± 1.04 cd	174.72 ± 19.66 d	176.70 ± 10.57 d
CC	29.41 ± 4.67 c***	5.39 ± 0.14 d	12.82 ± 1.18 c***	4.53 ± 0.51 e	205.17 ± 44.86 d***	72.26 ± 6.20 e
HS	32.17 ± 1.72 c***	10.51 ± 0.91 c	12.97 ± 1.52 c	12.95 ± 0.83 c	396.81 ± 27.27 c***	265.27 ± 10.21 c
MS	24.37 ± 1.25 d***	8.39 ± 0.44 c	9.06 ± 0.79 c***	6.70 ± 0.32 de	198.54 ± 29.20 d***	102.24 ± 5.14 e

¹ Different lowercase letters indicate significant differences within a column among TE, LJ, CA, CC, HS, and MS, as determined by one-way ANOVA (p < 0.05). Statistical significance within a row between hot-water and 80% ethanol is indicated as *** (p < 0.0001), * (p < 0.05) according to the t-test. ² Details of the sample code are provided in Figure 1.

presents the results of antioxidant activity analyses, including ABTS, DPPH, and FRAP assays and

Table 3. Pearson correlation ¹ between TPC, TFC, and four total antioxidant capacity (TAC) values of hot-water extracts and 80% ethanol extracts.

	TPC	TFC	ABTS	DPPH	FRAP
TPC	1	0.67 *	0.86 *	0.93 *	0.91 *
TFC	-	1	0.91 *	0.72 *	0.89 *
ABTS	-	-	1	0.82 *	0.95 *
DPPH	-	-	-	1	0.92 *
FRAP	-	-	-	-	1

¹ Values represent Pearson correlation coefficients (r) between TPC, TFC, and TAC (ABTS, DPPH, and FRAP). An asterisk (*) indicates a significant correlation at p < 0.05.

shows the correlations among antioxidant activities. Consistent with the higher TPC values, hot-water extracts exhibited stronger antioxidant capacities than 80% ethanol extracts across most EFTs.

The ABTS radical scavenging activity of hot-water extracts ranged from 101.41 ± 4.15 to 24.37 ± 1.25 mg vitamin C equivalent/g dw, showing significantly greater antioxidant potential than 80% ethanol extracts. TE displayed the strongest activity, while MS showed the weakest. Most other samples demonstrated superior radical scavenging performance in hot-water extracts, indicating matrix-dependent efficiency of hydrophilic antioxidants. The DPPH assay results further confirmed the superior antioxidant effect of hot-water extracts. The DPPH scavenging capacities of hot-water extracts ranged from 66.56 ± 4.71 mg VCE/g dw (LJ) to 9.06 ± 0.79 mg VCE/g dw (MS), which were consistently higher than those of the corresponding 80% ethanol extracts (57.08 ± 5.93 mg VCE/g dw to 4.53 ± 0.51 mg VCE/g dw). Similarly, in the FRAP assay, the hot-water extracts exhibited greater ferric ion reducing power than the 80% ethanol extracts for all EFTs except CA. The reducing capacity of hot-water extracts ranged from 1345.91 ± 118.43 μM FeSO₄/g dw (TE) to 174.72 ± 19.66 μM FeSO₄/g dw (CA). These findings confirm that hot-water infusion more effectively releases phenolic compounds with high reducing capacity, contributing to overall stronger antioxidant potential, compared to 80% ethanol extraction.

3.3. Identification of Phenolic Compounds in Hot-Water Extracts of EFTs

The phenolic compounds identified in the six EFTs using UPLC-ESI-Q-TOF-MS in both negative and positive ionization modes are listed in

Table 4. Identification of phenolic compounds in hot-water extracts of six dried edible flowers using UPLC-ESI-Q-TOF-MS in negative mode and positive mode.

Proposed Compounds	RT (min)	Mode	m/z	Major Fragments	Formula	Classification	Species
Protocatechuic acid 4-O-glucoside	3.87	[M-H]¯	315.07	141, 185	C13H16O9	Phenolic acid	HS
Neochlorogenic acid	4.43	[M-H]¯	353.09	191, 135, 179	C16H18O9	Phenolic acid	TE, LJ, CA, CC, HS, and MS
Chlorogenic acid	5.04	[M-H]¯	353.09	191, 133, 215	C16H18O9	Phenolic acid	TE, LJ, CA, CC, HS, and MS
(−)-Cryptochlorogenic acid	5.21	[M-H]¯	353.09	173, 135, 179	C16H18O9	Phenolic acid	TE, LJ, CA, CC, HS, and MS
m-Coumaric acid	5.26	[M-H]¯	163.04	119, 145, 117	C9H8O3	Phenolic acid	TE, LJ, CA, CC, HS, and MS
Myricetin 3-galactoside	6.22	[M-H]¯	479.08	317, 316, 315	C21H20O13	Flavonoid	TE, LJ, CA, HS, and MS
Taxifolin 3-O-rhamnoside	6.31	[M-H]¯	449.11	259, 287, 121	C21H22O11	Flavonoid	TE, LJ, CA, CC, HS, and MS
Quercetin 3-O-glucosyl-xyloside	6.34	[M-H]¯	595.13	371, 597, 335	C26H28O16	Flavonoid	HS
Quercetin 3-O-(6′″-trans-p-coumaroyl-2″-glucosyl)rhamnoside	6.35	[M-H]¯	755.2	284, 300, 255	C21H20O12	Flavonoid	CA
Rutin	6.64	[M-H]¯	609.15	300, 271, 255	C27H30O16	Flavonoid	TE, LJ, CA, CC, HS, and MS
Isoquercitrin	6.9	[M-H]¯	463.09	301, 285, 271	C21H20O12	Flavonoid	TE, LJ, CA, CC, and HS
Luteolin 7-O-glucoside	6.97	[M-H]¯	447.09	285, 113	C21H20O11	Flavonoid	TE, LJ, HS, and MS
Kaempferol 3-O-rutinoside	7.16	[M-H]¯	593.15	285, 284, 255	C27H30O15	Flavonoid	TE, LJ, CA, CC, HS, and MS
3,4-Dicaffeoylquinic acid	7.38	[M-H]¯	515.12	353, 179, 135	C25H24O12	Phenolic acid	CA, CC, and HS
Genistein 7-O-glucuronide	7.66	[M-H]¯	445.09	269, 268, 431	C21H18O11	Flavonoid	TE, LJ, and CC
Kaempferol 3-O-glucoside	7.4	[M-H]¯	447.09	191, 255, 255	C21H20O11	Flavonoid	LJ
3,5-Di-O-caffeoylquinic acid	7.71	[M-H]¯	515.12	353, 179, 135	C25H24O12	Phenolic acid	TE, LJ, CA, CC, HS, and MS
Kaempferol 3,7-O-diglucoside	4.23	[M+H]+	611.16	287, 449, 137	C57H110O16Si10	Flavonoid	CC and MS
Quercetin 3-sambubioside	4.56	[M+H]+	597.14	303, 301	C26H28O16	Flavonoid	HS
Kaempferol 3-O-xylosyl-glucoside	4.96	[M+H]+	581.15	287, 549, 137	C15H10O6	Flavonoid	HS
Kaempferol 3-O-α-L-[6′″-p-coumaroyl-β-D-glucopyranosyl-(1→2)-rhamnopyranoside]-7-O-β-D-glucopyranoside	5.96	[M+H]+	903	271, 287, 609	C42H46O22	Flavonoid	LJ
Quercetin 3-neohesperidoside-7-rhamnoside	6.34	[M+H]+	757.22	287, 303, 153	C33H40O20	Flavonoid	CA
Quercetin	6.87	[M+H]+	303.05	273, 153, 121	C15H10O7	Flavonoid	TE, LJ, CA, CC, HS, and MS
Luteolin 7-O-rhamnoside	6.92	[M+H]+	595.16	287, 449, 117	C27H30O15	Flavonoid	TE, LJ, CA, CC, HS, and MS
Herbacetin	8.41	[M+H]+	303.05	169, 121	C15H10O7	Flavonoid	TE, CA, CC, and MS

, which also displays the heatmap of the phenolic compounds in the hot-water extracts (Figure 2). In negative ion mode, several phenolic acids—such as neochlorogenic acid, chlorogenic acid, (−)-cryptochlorogenic acid, m-coumaric acid, and 3,5-di-O-caffeoylquinic acid—were detected in all samples. Flavonoids including taxifolin 3-O-rhamnoside, rutin, and kaempferol 3-O-rutinoside were also identified in all EFTs. Myricetin 3-galactoside was detected in five species except CC, while kaempferol 3-O-glucoside and quercetin 3-O-(6′″-trans-p-coumaroyl-2″-glucosyl)rhamnoside were specifically observed in LJ and CA, respectively. Protocatechuic acid 4-O-glucoside and quercetin 3-O-glucosyl-xyloside were unique to HS. Among flavonoids, isoquercitrin was detected in all EFTs except MS, luteolin 7-O-glucoside was detected in all except CA and CC, and genistein 7-O-glucuronide was found in TE, LJ, and CC. 3,4-Dicaffeoylquinic acid was identified in CA, CC, and HS. In positive ion mode, quercetin and luteolin 7-O-rhamnoside were detected in all EFTs. Herbacetin was found in all samples except LJ and HS, while kaempferol 3,7-O-diglucoside appeared in CC and MS. Quercetin 3-sambubioside, kaempferol 3-O-xylosyl-glucoside, and quercetin 3-O-neohesperidoside-7-rhamnoside were uniquely found in HS, and kaempferol 3-O-α-L-[6′″-p-coumaroyl-β-D-glucopyranosyl-(1→2)-rhamnopyranoside]-7-O-β-D-glucopyranoside was identified in LJ.

3.4. Identification of Volatile Organic Compounds in Hot-Water Extracts of EFTs

Table 5. Identification of volatile organic compounds in six dried edible flower petals using HS-SPME/GC-MSR.

Sample Code 1	RT (min)	m/z (Major Fragment Ion)	Formula	Classification	Proposed Compounds	Odor
TE	7.21	72, 40, 41	C4H8O	Aldehyde	Isobutyraldehyde	pungent
	8.33	86, 43, 42	C4H6O2	Ketone	2,3-Butanedione	butter
	12.37	86, 44, 41	C5H10O	Aldehyde	3-Methylbutanal	fruity
	12.99	96, 57, 41	C5H12O	Aldehyde	2-Methylbutanal	sweet
	18.39	88, 42, 55	C5H12O	Alcohol	1-Pentanol	sweet
	19.50	100, 44, 56	C6H12O	Aldehyde	Hexanal	green grass
	21.35	128, 43, 85	C9H20	Alkane	2,4-Dimethylheptane	pungent
	26.23	136, 93, 91	C10H16	Monoterpene	α-pinene	woody
	28.00	136, 93, 77	C10H16	Monoterpene	Sabinene	citrus; woody
	28.54	136, 93, 41	C10H16	Monoterpene	Myrcene	floral
	30.55	136, 121, 93	C10H16	Monoterpene	α-terpinene	woody; mint
	31.21	136, 93, 91	C10H16	Monoterpene	(E)-β-Ocimene	green grass
	31.37	136, 68, 93	C10H16	Monoterpene	D-Limonene	citrus
	31.93	136, 93, 91	C10H16	Monoterpene	(Z)-β-ocimene	green grass
	32.80	136, 93, 121	C10H16	Monoterpene	gamma-Terpinene	woody
	34.21	136, 93, 121	C10H16	Monoterpene	Terpinolene	woody
	39.88	152, 82, 110	C10H16O	Monoterpene ketone	Piperitone	mint
	43.37	150, 107, 91	C10H14O	Monoterpene ketone	Piperitenone	mint
	47.07	204, 93, 91	C15H24	Sesquiterpene	(E)-Caryophyllene	spicy
LJ	5.35	58, 29, 28	C3H6O	Aldehyde	Propanal	fruity
	6.41	74, 73, 42	C3H6O2	Ester	Methyl Acetate	fruity
	7.22	72, 40, 41	C4H8O	Aldehyde	Isobutyraldehyde	pungent
	7.66	70, 41, 39	C4H6O	Aldehyde	Methacrolein	pungent
	8.35	86, 43, 42	C4H6O2	Ketone	2,3-Butanedione	butter
	8.47	86, 43, 42	C6H14	Alkane	2-Methylpentane	gasoline
	11.69	70, 41, 39	C4H6O	Aldehyde	2-Butenal	alcoholic
	12.38	86, 44, 41	C5H10O	Aldehyde	3-Methylbutanal	fruity
	12.99	96, 57, 41	C5H12O	Aldehyde	2-Methylbutanal	sweet
	14.06	84, 56, 27	C5H8O	Ketone	1-Penten-3-one	spicy; pungent
	14.32	86, 57, 29	C5H10O	Alcohol	1-Penten-3-ol	burnt; butter
	14.55	100, 43, 29	C5H8O2	Ketone	2,3-Pentanedione	butter
	14.65	86, 44, 29	C5H10O	Aldehyde	Pentanal	green grass; fruity; pungent
	15.49	96, 81, 53	C6H8O	Heterocyclic aromatic compound	2-Ethylfuran	pungent; burnt
	17.31	84, 55, 83	C5H8O	Aldehyde	trans-2-Pentenal	fruity
	18.41	88, 42, 55	C5H12O	Alcohol	1-Pentanol	sweet
	19.50	100, 44, 56	C6H12O	Aldehyde	Hexanal	green grass
	20.43	184, 43, 85	C13H28	Alkane	2,4-Dimethylhexane	citrus; green grass; woody
	21.35	128, 43, 85	C9H20	Alkane	2,4-Dimethylheptane	pungent
	21.44	114, 41, 55	C7H14O	Aldehyde	trans-2-Hexenal	almond
	21.88	100, 67, 41	C6H12O	Alcohol	3-Hexenol	green grass
	22.38	102, 56, 55	C6H14O	Alcohol	1-Hexanol	sweet; green grass
	24.62	130, 74, 43	C7H14O2	Ester	Methyl hexanoate	fruity
	26.05	106, 77, 105	C7H6O	Aldehyde	Benzaldehyde	almond
	30.56	204, 57, 43	C12H25C	Alkane	2,5-Dimethylnonane	citrus
	31.09	184, 71, 43	C13H28	Alkane	4,6-Dimethyldodecane	floral; sweet
	33.11	184, 71, 57	C13H28	Alkane	5-(2-Methylpropyl)nonane	gasoline
CA	6.23	94, 62, 47	C2H6S	Sulfur compound	Dimethyl sulfide	sweet
	6.39	74, 73, 42	C3H6O2	Ester	Methyl Acetate	fruity
	7.22	72, 40, 41	C4H8O	Aldehyde	Isobutyraldehyde	pungent
	12.38	86, 44, 41	C5H10O	Aldehyde	3-Methylbutanal	butter
	13.00	96, 57, 41	C5H12O	Aldehyde	2-Methylbutanal	fruity
	14.65	86, 44, 29	C5H10O	Aldehyde	Pentanal	green grass; fruity; pungent
	16.97	88, 55, 42	C5H12O	Alcohol	Isoamyl Alcohol	alcoholic
	19.50	100, 44, 56	C6H12O	Aldehyde	Hexanal	green grass
	21.36	128, 43, 85	C9H20	Alkane	2,4-Dimethylheptane	pungent
	22.95	128, 43, 85	C9H20	Alkane	4-Methyloctane	pungent
	31.09	142, 71, 57	C10H22	Alkane	4,6-Dimethyldodecane	floral; sweet
	33.12	184, 71, 57	C13H28	Alkane	5-(2-Methylpropyl)nonane	gasoline
CC	6.40	74, 73, 42	C3H6O2	Ester	Methyl Acetate	fruity
	7.22	72, 40, 41	C4H8O	Aldehyde	Isobutyraldehyde	pungent
	8.34	86, 43, 42	C4H6O2	Ketone	2,3-Butanedione	butter
	12.38	86, 44, 41	C5H10O	Aldehyde	3-Methylbutanal	fruity
	13.00	96, 57, 41	C5H12O	Aldehyde	2-Methylbutanal	sweet
	14.33	86, 57, 29	C5H10O	Alcohol	1-Penten-3-ol	burnt; butter
	14.65	86, 44, 29	C5H10O	Aldehyde	Pentanal	green grass; fruity; pungent
	18.41	88, 42, 55	C5H12O	Alcohol	1-Pentanol	sweet
	19.50	100, 44, 56	C6H12O	Aldehyde	Hexanal	green grass
	21.36	128, 43, 85	C9H20	Alkane	2,4-Dimethylheptane	pungent
	29.50	156, 57, 43	C11H24	Alkane	Undecane	faint
	33.12	184, 71, 57	C13H28	Alkane	5-(2-Methylpropyl)nonane	gasoline
HS	6.38	74, 73, 42	C3H6O2	Ester	Methyl Acetate	fruity
	19.51	100, 44, 56	C6H12O	Aldehyde	Hexanal	green grass
	20.44	74, 73, 42	C3H6O2	Aldehyde	Furfural	sweet; almond
MS	8.35	86, 43, 42	C4H6O2	Ketone	2,3-Butanedione	butter
	12.39	86, 44, 41	C5H10O	Aldehyde	3-Methylbutanal	fruity
	13.00	96, 57, 41	C5H12O	Aldehyde	2-Methylbutanal	sweet
	14.33	86, 57, 29	C5H10O	Alcohol	1-Penten-3-ol	burnt; butter
	14.66	86, 44, 29	C5H10O	Aldehyde	Pentanal	green grass; fruity; pungent
	18.41	98, 69, 41	C6H10O	Ketone	Isopropenyl ethyl ketone	pungent
	19.51	100, 44, 56	C6H12O	Aldehyde	Hexanal	green grass
	21.36	128, 43, 85	C9H20	Alkane	2,4-Dimethylheptane	pungent
	30.23	138, 67, 81	C10H18	Alkyne	1-Decyne	green grass; citrus
	32.06	124, 54, 67	C9H16	Alkene	1,2-Nonadiene	gasoline
	33.13	184, 71, 57	C13H28	Alkane	5-(2-Methylpropyl)nonane	gasoline

¹ Details of the sample code are provided in Figure 1.

summarizes the qualitative analysis of VOCs in EFTs using HS-SPME/GC-MS, which also displays the heatmap of VOCs (Figure 3). A total of 4, 11, 5, 5, 2, and 4 aldehydes were identified in TE, LJ, CA, CC, HS, and MS, respectively. Hexanal was detected in all samples, and isobutyraldehyde was detected in TE, LJ, CA, and CC. 3-Methylbutanal and 2-methylbutanal were detected in five samples except HS, and pentanal was observed in LJ, CA, CC, and MS. Specifically, propanal, methacrolein, 2-butenal, trans-2-pentenal, trans-2-hexenal, and benzaldehyde were additionally detected in LJ, and furfural was detected in HS. The number of ester compounds detected was two in LJ, one in CA, one in CC, and one in HS. Methyl acetate was detected in all four samples (LJ, CA, CC, and HS), while methyl hexanoate was detected only in LJ. The numbers of ketone compounds detected were one in TE, three in LJ, one in CC, and two in MS. 2,3-Butanedione was found in all four EFTs (TE, LJ, CC, and MS), while 1-penten-3-one and 2,3-pentanedione were specifically detected in LJ, and isopropenyl ethyl ketone was specifically detected in MS. The numbers of alcohol compounds detected were one in TE, four in LJ, one in CA, two in CC, and one in MS, while none were detected in HS. 1-Pentanol was found in TE, LJ, and CC, and 1-penten-3-ol was detected in LJ, CC, and MS. Specifically, 3-hexenol and 1-hexanol were identified in LJ, and isoamyl alcohol was identified in CA. The numbers of alkane compounds identified were one in TE, six in LJ, four in CA, three in CC, and two in MS. Among the five samples in which alkanes were present, 2,4-dimethylheptane was detected in all of them. In contrast, 5-(2-methylpropyl)nonane was found in the same five samples except TE, and 4,6-dimethyldodecane was identified only in LJ and CA. Specifically, 2-methylpentane, 2,4-dimethylhexane, and 2,5-dimethylnonane were detected in LJ, 4-methyloctane in CA, and undecane in CC. In addition to the major compounds mentioned above, there are compounds detected solely in specific samples. Monoterpenes were detected exclusively in TE, including α-pinene, sabinene, myrcene, α-terpinene, (E)-β-ocimene, D-limonene, (Z)-β-ocimene, gamma-terpinene, terpinolene, piperitone, and piperitenone. The sesquiterpene (E)-caryophyllene was also uniquely identified in TE. At the same time, the heterocyclic aromatic compound 2-ethylfuran was detected in LJ, the sulfur compound dimethyl sulfide was detected in CA, and the alkyne 1-decyne and the alkene 1,2-nonadiene were detected in MS.

4. Discussion

To examine the efficacy of hot-water extract to consume edible flowers, this study used six EFTs available in Korea to compare hot-water and 80% ethanol extractions. As a result, significant differences were observed across all analyses between species and ex-traction solvents. A key finding was the significant impact of solvent polarity on extraction yield. For TPC, hot-water extraction proved superior, yielding results 1.3 to 4 times higher than 80% ethanol. This trend is consistent with Amensour et al. [23], who observed higher TPC from Myrtus communis L. using hot water compared with room-temperature ethanol. The superior performance of hot water can be attributed to its high polarity, which promotes solubilization of hydrophilic acids [23,24]. Phenolic acids, characterized by a benzene ring with attached carboxyl and hydroxyl groups, are inherently polar and thus dissolve readily in water.

However, the previous literature indicates that the relative efficiency of water versus ethanol for TPC is not universal and can shift depending on extraction temperature and plant species. Studies comparing the two solvents at identical temperatures—whether at room temperatures and moderate temperatures (e.g., 50 °C)—often report higher TPC yields with hydroethanolic solvents (30−70%). Likewise, Suksathan et al. [25] demonstrated that when boiling water was compared with 95% ethanol, the superior solvent differed by species, with some flowers exhibiting higher TPC in water and others showing higher yields in ethanol. These observations highlight that TPC extraction is jointly governed by solvent polarity, extraction temperature, and species-specific phytochemical composition. Within this context, the present findings suggest that under consumer-relevant hot-water infusion conditions, water remains highly effective for recovering phenolic acids from edible flower petals.

In contrast, the TFC values of two extraction methods were largely similar in magnitude, despite ethanol yielding higher values in several samples. It suggested that both extraction methods were similarly effective in recovering flavonoid compounds, despite ethanol being well known as the most commonly used solvent for flavonoid extraction with a high efficiency [11]. This difference between TPC and TFC stems from the fundamental structural characteristics between phenolic acids and flavonoids, whose larger backbone structure of two benzene rings and a heterocyclic ring confers a generally lower polarity compared to the smaller, more hydrophilic phenolic acids. Consequently, flavonoids tend to dissolve more effectively in solvents of intermediate polarity, such as 80% ethanol [9]. Previous studies also support this behavior. For example, hydroalcoholic system has been reported to yield higher flavonoid recovery than water, reflecting the polarity–solubility match of flavonoid compounds [26,27]. However, Suksathan et al. [25] demonstrated that TFC differed by species when comparing boiling water and 95% ethanol. Such species-dependent variability provides an explanation for why the TFC patterns in this study also differed among the six edible flowers. Nevertheless, our results confirm that hot-water extraction, mimicking the preparation of tea for consumption, is still capable of extracting a substantial amount of these beneficial compounds.

As discussed above, the high temperature of the hot-water extraction also played a crucial role in enhancing the yield of phenolic compounds. Heat can disrupt plant cell wall structures, breaking the bonds between lignin and phenolic acids and thereby re-leasing otherwise insoluble phenolics [28]. Furthermore, thermal degradation of lignin it-self can generate new free phenolic acids, further increasing the total measured content. This thermal effect, as described by Kim et al. [29] in their study on heat-treated Oryza sativa L., helps explain the high TPC observed in the hot-water extracts, where the TAC results strongly correlated with the TPC and TFC values [29], as also confirmed in Table 3 of this study. This correlation confirms that the phenolic compounds—encompassing both phenolic acids and flavonoids—serve as the main factors driving the extracts’ antioxidant properties, namely their free-radical scavenging ability and reducing power. Among the edible flowers tested, TE consistently exhibited the highest TAC. This indicates that TE possesses a superior quantitative and qualitative composition of these bioactive phenolic compounds. These findings are consistent with the work of Dujmović et al. [10], who also reported that the TPC and TFC of Tagetes erecta L. were 1.4 to 2 times higher than those of cornflower and common marigold, establishing a strong positive correlation between phenolic composition and antioxidant capacity.

In this study, hot-water extraction was adopted to mimic the actual consumption conditions of edible flower teas, whereas 80% ethanol extraction served as an analytical reference commonly used in industrial and laboratory settings for isolating functional compounds. Therefore, the methodological differences between hot-water and ethanol extraction in this study (e.g., concentration of flower petal and solvent, temperature, and physical treatment) should be considered when interpreting these results. Because hot-water extraction more accurately reflects real-world ingestion patterns and was the primary focus of evaluating bioactive components under consumer-relevant conditions, the detailed profiling of phenolic compounds was subsequently conducted on only the hot-water extracts using UPLC-ESI-Q-TOF-MS.

The analysis revealed distinct differences across six EFTs in the types and levels of major phenolic acids and flavonoids detected across samples. This suggests that even with the same extraction method, the variety and metabolic characteristics of the raw material have a significant impact on the component extraction efficiency and composition. The Table 4 analysis results show that most samples were relatively rich in phenolic acids such as caffeic acid, chlorogenic acid, and coumaric acid, and flavonoids such as rutin, quercetin, and luteolin glycosides were predominantly detected. The reason for these results is that the structure in which flavonoids and glycosides are combined has high polarity, making it easy to dissolve during hot-water extraction. Glycosides of flavonoid compounds increase their solubility and stability in water, increase their bioavailability, and reduce their potentially harmful bioactive effects on the body [30]. In particular, EFTs rich in flavonoids are likely to have stronger antioxidants and anti-inflammatory activities, as reported in previous studies [3,5,16,25], while samples high in phenolic acids are likely to contribute not only to antioxidant activity but also to antibacterial activity.

GC–MS analysis revealed that the six edible flower petal teas exhibited distinct VOCs profiles characterized by monoterpenes (α-pinene, (E)/(Z)-β-ocimene, and terpinolene), aldehydes (isobutyraldehyde, hexanal, and pentanal), ketones (2,3-butanedione and isopropenyl ethyl ketone), esters (methyl acetate and methyl hexanoate), and alcohols (1-Pentanol, 1-Penten-3-ol, and 1-Hexanol). These compounds are readily extractable into the aqueous phase, consistent with the findings of Slámová et al. [30], which demonstrated that polar VOCs preferentially partition into the water phase, indicating that water-infused EFTs can effectively release characteristic aromatic compounds. From a sensory perspective, the compositional differences identified by GC–MS correspond closely to the characteristic aroma notes perceived in each flower petal tea. Monoterpenes such as α-pinene, D-limonene, and β-ocimene contribute woody, citrus, and herbal top notes, imparting freshness and brightness reminiscent of green or herbal teas [31]. Aldehydes including hexanal and pentanal reinforce this green, grassy freshness, which enhances the perception of naturalness [32]. Ketones such as 2,3-butanedione and isopropenyl ethyl ketone provide buttery and creamy undertones, softening the aroma profile, while esters (methyl acetate and methyl hexanoate) add sweet and fruity nuances that increase consumer preference [33,34]. Alcohols (1-hexanol and isoamyl alcohol) further enrich floral sweetness, and furan derivatives (furfural and 2-ethylfuran) contribute roasted, caramel-like base-notes that deepen the overall flavor complexity [35].

The sensory contribution of individual VOCs is influenced not solely by their concentration but also by their odor thresholds and synergistic interactions. For instance, hex-anal—detected across all EFTs samples—can evoke earthy, musty odors at 10–100 ppm but transitions to a green, grassy note at higher levels [36]. Thus, the interplay among compounds with contrasting thresholds generates a multilayered aromatic perception. Taken together, the GC–MS data demonstrate that the aromatic identity of each flower tea is defined by its specific metabolic VOC fingerprint, where the relative abundance of terpenes, aldehydes, alcohols, and esters shapes the sensory experience. The distinctive VOC compositions observed across flower species highlight species-specific metabolic pathways governing aroma biosynthesis and the resulting sensory individuality when consumed as tea.

5. Conclusions

This study demonstrates that hot-water infusion is an efficient and sustainable method for extracting functional and aromatic compounds from edible flower petals. Compared with 80% ethanol, hot-water extraction yielded significantly higher TPCs and antioxidant activities (ABTS, DPPH, and FRAP), confirming that hydrophilic phenolic acids are preferentially solubilized in aqueous media. UPLC–ESI–Q-TOF–MS and GC–MS analyses revealed that hot-water extracts contained abundant chlorogenic acid derivatives, flavonoid glycosides, and aroma compounds such as α-pinene and hexanal, defining the sensory characteristics of flower teas.

These findings establish a scientific basis for promoting flower teas as natural, safe, and functional beverages. Furthermore, from an industrial perspective, hot-water extraction offers some advantages in terms of safety and sustainability. It requires no organic solvents, thereby reducing both environmental impact and processing costs. It also aligns with the recent trend toward pollution-free extraction technologies (green extraction technologies), which emphasize low-toxicity or solvent-free methods for recovering these active compounds. These attributes make hot-water extraction a promising method for large-scale, eco-friendly production of functional ingredients.