Comparative evaluation of bioactive compounds and antioxidant activity in Justicia spicigera extracts using ultrasound-assisted and microwave-assisted extraction methods

Abstract

A comparative study of ultrasound-assisted extraction (UAE) was conducted, and for the first time, microwave-assisted extraction (MAE) was reported for the recovery of bioactive compounds from Justicia spicigera. Among the compounds identified in MAE were kaempferol and peonidin derivatives. In contrast, the extracts obtained via UAE contained cyanidin and kaempferol derivatives. These compounds were associated with the colorimetric changes observed as a function of pH. The results demonstrated that MAE achieved a higher recovery of phenolic compounds (48.93 mg GAE/g of dry extract) and total flavonoids (23.46 mg catechin/g of dry extract). Regarding antioxidant activity, MAE exhibited an IC₅₀ of 124.47 μg of dry extract/mL in the DPPH assay, with no significant differences compared to ABTS and FRAP assays when compared to UAE. This study underscores the role of both extraction technologies in the recovery of bioactive compounds from Justicia spicigera and the potential development of active and intelligent packaging.

Introduction

Justicia spicigera (J. spicigera), a plant native to Mexico and South America, is commonly known as Muicle or Muitle (Anaya-Esparza et al., 2018). The main phytochemicals reported in this species include kaempferol derivatives (kaempferol-trirhamnoside). In addition, various anthocyanins have been identified, such as cyanidin 3-O-glucoside and cyanidin 3,5-diglucoside. Currently, research on the effects of J. spicigera is on the treatment of various diseases (Anaya-Esparza et al., 2018). Another notable property of J. spicigera is its dyeing capacity. J. spicigera has been used to dye textiles and handicrafts, and recently, they have been incorporated as a natural colorant in various food products (Baqueiro-Peña and Guerrero-Beltrán, 2017).

Traditionally, the recovery of bioactive compounds has been performed using conventional techniques such as maceration, organic solvent extraction, and Soxhlet extraction, among others. However, despite being efficient for compound recovery, these techniques require prolonged extraction times and large amounts of solvents and may lead to the thermal degradation of heat-sensitive compounds (Platzer et al., 2022). Studies have been conducted to address these challenges and develop new extraction methods that reduce extraction times, solvent consumption, energy requirements, and costs to counteract the main disadvantages of using conventional techniques. These methods are ultrasound-assisted extraction (UAE) and microwave-assisted extraction (MAE). Since phytochemicals are typically found within cells and surrounded by a rigid cell wall, employing appropriate and efficient extraction technologies is essential (Imtiaz et al., 2023).

The principle of UAE is based on cavitation, which involves the formation, growth, and collapse of vapor or gas bubbles generated by ultrasonic waves at a specific frequency. This process disrupts the cell wall, allowing the solvent to penetrate the plant material and facilitating the release of the target compounds (Shen et al., 2023). Another emerging technology for the recovery of bioactive compounds is MAE, which operates through the energy supplied by the dissipation of electromagnetic waves. These waves induce molecular movement and friction within the medium, generating electroporation and vaporization of the intracellular water. These effects lead to increased pressure, causing the rupture of cell walls and facilitating the release of target molecules (Walayat et al., 2024). Although the effects of UAE on the recovery of bioactive compounds from J. spicigera have been previously demonstrated (Baqueiro-Peña and Guerrero-Beltrán, 2017; Hernández-Rodríguez et al., 2020), it is relevant to compare this technology with an emerging extraction technique that employs a different mechanism, such as MAE, which has not yet been reported for the extraction of bioactive compounds from this species.

An alternative to valorize J. spicigera extract could be its application in developing active and intelligent packaging. The development of active packaging requires extracts with a high concentration of bioactive compounds for their functionality. On the other hand, intelligent packaging requires specific compounds to monitor product freshness. Due to their operating principles, emerging extraction technologies such as UAE and MAE can facilitate the recovery of bioactive compounds and simultaneously extract compounds with different polarities, sizes, or chemical structures. Improvement of quantity and quality of compounds could contribute to the application of these extracts in the development of active and intelligent packaging (Ahmed et al., 2022; Sarkar et al., 2025).

This study aimed to extract bioactive compounds from J. spicigera using UAE and MAE. The bioactive compounds obtained were identified by high-performance liquid chromatography. In the second stage, their physicochemical characteristics were evaluated using stability tests, color changes, and quantification of phenolic compounds, flavonoids, and antioxidant capacity.

Materials and methods

Plant material

The aerial parts (stems and fresh leaves) of J. spicigera were collected in Texcoco de Mora, State of Mexico, Mexico (19°29′23″N, 98°53′37″W). Taxonomic identification was performed in the Herbarium “JES” of the Universidad Autónoma Chapingo. The collected plant material was washed and dried in an oven (Fe-294AD, Feligneo, Jalisco-Mexico) at 30 ± 5 °C for 3 days. Subsequently, the dry sample was pulverized in a disk mill (GX4100, Krups, Germany) and sieved through a mesh with an opening of 841 μm. Finally, the dry powder was stored in plastic bags in dark conditions and at room temperature until use.

Extractions

Two extraction methods were used in this study: ultrasound assisted extraction and microwave assisted extraction. The methodology described by Vargas-Torrico et al. (2022) was used to perform extractions. As extraction solvent, a mixture of ethanol and water was used in a 70:30 (v/v) ratio, using a ratio of 1:20 (g sample/mL solvent).

Ultrasound assisted extraction (UAE)

The plant material was allowed to stand with the solvent for 30 min at room temperature (28–30 °C). Subsequently, the mixture was sonicated in an ultrasonic bath (TI-H-5, Elma, Germany) at a frequency of 45 kHz for 30 min, at 65 ± 2 °C. The extracts were centrifuged (model K, International Equipment Co., USA) at 1750 rpm for 15 min and filtered with Whatman No. 1 paper (Whatman® International, Ltd., England). Finally, the filtered extracts were concentrated at a rotary evaporator (RE-500, Yamato, Japan) at 35 ± 2 °C and freeze-dried (FreeZone 4.5, Labconco™, USA). The extracts were stored in amber vials at room temperature until use.

Microwave assisted extraction (MAE)

The plant material was left to stand for 30 min at room temperature (28–30 °C). The mixture was placed in a microwave oven (Labtron LMWD-A10, UK) at 30 psi pressure for 15 min at 65 ± 2 °C. Centrifugation, lyophilization and storage conditions were maintained as described in UAE methodology.

Identification of compounds by LC–QTOF MS analysis

The determination of the chemical components in each of the extracts was carried out by high performance liquid chromatography (HPLC) coupled to mass spectroscopy (ESI-QTOF), using an HPLC-QTOF system (Agilent Model G6530, Agilent Technologies, Palo Alto, CA, USA). This equipment is composed with a Poroshell 120 EC- C8 column (50 × 3.0 mm, 2.7 µm particle size), was maintained at a temperature of 30 °C. The mobile phases consisted of (A) 0.1% water with formic acid and (B) acetonitrile. Compound separation was carried out with the following elution gradient: initial conditions 5% B; 5 min, 10% B; 7 min, 15% B; 10 min, 20% B; 12 min, 25% B; 15 min, 55% B; 17 min, 85% B; 20 min, 5% B; with a flow rate of 0.4 mL/min and an injection volume of 7.0 µL.

Compound identification was carried out using MassHunter Software (Agilent, Santa Clara, CA, USA) and mass spectrometry databases, such as The Human Metabolome Database (www.hmdb.ca).

Stability of extracts

Sensitivity to pH and UV–Vis spectra

For the pH sensitivity analysis, the methodology described by Jiménez-González et al. (2023) with some modifications. Five mg of lyophilized extract was weighed and dissolved in 5 mL of distilled water adjusted to different pH values (2–12). The pH of each solution was adjusted by adding hydrochloric acid (HCl) or sodium hydroxide (NaOH) as needed. The solutions were stirred for 1 min to ensure homogeneous mixing. Subsequently, the absorption spectra were recorded in the range 200 to 800 nm using a Lambda 35 spectrophotometer (Perkin Elmer Ltd., USA) to obtain the absorption profile of each solution as a function of pH.

Color analysis

The various extract added with buffers described in the pH sensitivity analysis were measured using a high-resolution colorimeter (NH300, China) using a CIE Lab scale. For each measurement, the values of a* (greenish/reddish) and b* (blue/yellowish) were recorded to calculate the hue angle (Hue) according to Eq. (1):

$$Hue={\mathit{tan}}^{-1}(b_{}^{\ast }/a_{}^{\ast })$$ 1

The total color difference (ΔE) was calculated using Eq. (2):

$$\mathrm{\Delta }\text{E}=\sqrt{(L_{}^{\ast }-L_0^{\ast })_{}^2+(a_{}^{\ast }-a_0^{\ast })_{}^2+(b_{}^{\ast }-b_0^{\ast })_{}^2}$$ 2

where $L_0^{\ast }$, $a_0^{\ast }andb_0^{\ast }$ are the color parameters at time zero (pH 2) and L*, a* and b* are the color parameters at any time (Jiménez-González et al., 2023).

Phytochemical composition

Determination of total phenolic compounds (TPC)

The total phenolic content was calculated following the methodology of Singleton & Rossi (1965), with some modifications, using gallic acid as a standard. To a sample volume of 25 µL, 125 µL of distilled water was added, followed by 20 µL of Folin-Ciocalteu's reagent (1:10) and 30 µL of a 20% (m/v) sodium carbonate solution. After 30 min of resting in the dark, the absorbance was measured at 760 nm using a Multiskan Go (Thermo Fisher, Finland). The results were expressed as mg gallic acid equivalent per gram of dry extract (mg GAE/g dry extract).

Determination of total flavonoid compounds (TFC)

The total flavonoid content of the extracts was determined by a colorimetric procedure using aluminum chloride (AlCI₃) and sodium hydroxide (NaOH) (Kubola and Siriamornpun, 2011). In Falcon tubes, 0.5 mL of extract was mixed with 2.5 mL of distilled water and 0.15 mL of 5% sodium nitrite (NaNO₂) solution, the mixture was allowed to stand for 6 min. Then, 300 µL of 10% AlCl₃∙6H₂O was added and allowed to stand for 5 min, followed by adding 1 mL of 5% NaOH. Finally, the mixture was vortexed (3000 rpm) and the absorbance was measured at 510 nm using a Multiskan Go (Thermo Fisher, Finland). The results were expressed as mg catechin per gram of dry extract (mg catechin/g dry extract).

Determination of antioxidant activity

Free radical scavenging capacity (DPPH)

The antiradical capacity was determined by the DPPH (2, 2-diphenyl-1-picrylhydrazyl) assay. A 50 µL of a 1 mM DPPH solution was added to a 200 µL aliquot of extract (Brand-Williams et al., 1995). After 30 min, the absorbance was measured at 515 nm. The control consisted of 200 µL of 80% methanol with 50 µL of DPPH solution, while the blank consisted of 250 µL of 80% methanol. The percent reduction of DPPH was calculated using Eq. (3):

$$DPPHScaveningActivity(\%)=[1-(Am-Ab)/(Ac-Ab)]\times 100$$ 3

where Am is the absorbance of the sample, Ab is the absorbance of the blank, and Ac is the absorbance of the control.

The IC₅₀ value was calculated from the graph relating the percentage of DPPH degraded to the sample concentration. Antioxidant capacity was expressed as μg of dry extract per mL (μg of dry extract/mL).

Determination of ferric reducing antioxidant power (FRAP)

Antioxidant capacity was determined using the method of Benzie & Strain (1999). FRAP solution was prepared by mixing 10 mL of 300 mM acetate buffer (pH 3.6), 1 mL of TPTZ solution (0.01 M) and 1 mL of FeCl₆H₂0 solution (0.02 M). A 20 µL aliquot of extract was mixed with 180 µL of FRAP solution and allowed to stand for 30 min in the dark. The absorbance was read at 593 nm. The results were expressed as µM Trolox per gram of dry extract (µM Trolox /mg dry extract).

ABTS^•+ method: measurement of total antioxidant capacity (TAC)

The measurement of TAC by uptake of the ABTS radical was performed according to the methodology proposed by Miller & Rice-Evans (1996) with some modifications. Two solutions were prepared: a 7.4 mM ABTS solution and a 2.6 mM K₂S₂O₈ solution, which were combined in a 1:1 ratio. The mixture was allowed to stand in the dark for 16 h. Subsequently, it was diluted with methanol to an absorbance of 0.7—1.2 at 734 nm. To a sample volume of 20 µL, 180 µL of ABTS solution was added and allowed to stand for 10 min before reading in the spectrophotometer. The results were expressed as µM Trolox per mg of dry extract (µM Trolox/mg dry extract).

Statistical analysis

All results were expressed as the mean ± standard deviation. A completely randomized design was employed, and means were compared using Tukey’s test (α = 0.05). Statistical analyses were performed using SAS software (Statistical Analysis System, version 9.0 for Windows).

Results and discussion

Identification of compounds by LC–QTOF MS analysis

For the analysis and identification of the various compounds present in each extract, the molecular ion weight of each signal detected in the chromatogram and its fragments were considered (Yang et al., 2023). Additionally, compounds reported in the consulted literature were compared. The results of high-performance liquid chromatography (HPLC) coupled to electrospray ionization quadrupole time-of-flight mass spectrometry (ESI-QTOF) for the extracts obtained via ultrasound-assisted extraction (UAE) and microwave-assisted extraction (MAE) are presented in Tables 1 and 2.

Table 1.

Identification and classification of bioactive compounds in extracts of Justicia spicigera obtained by ultrasound assisted extraction using HPLC-DAD-ESI-QTOF mass coupled chromatography

TR (min)	Formula	Theoretical mass	[M + H]+ (m/z) cal	ESI–MS/MS Fragments (m/z)	Tentative identification	Type of compound
0.94	C22H18O10	442.089	442.37	118.0908, 315.1290	Epicatechin gallate	Flavanol
1.52	C41H43O17	807.249	807.235	286.1023, 625.1158, 404.1225	Malvidin 3- glucoside-ethyl-catechin	Anthocyanin
*2.38	C32H38O18	711.4	711.236	565.1745, 449.1226, 259.1736	Kaempferol-3-rhamnoside-7-xylosido-rhamnoside	Flavonol glycoside
*2.81	C33H40O18	724.7	725.252	579.1902, 433.1272, 242.1831	Kaempferol-3-rhamnoside-4''-rhamnoside-7-rhamnoside	Flavonoid
3.89	C40H41O18	809.229	809.281	621.2035, 433.1282, 234.1205	Malvidin 3- glucoside-ethyl-catechin	Anthocyanin
7.03	C32H31O14	639.171	639.407	309.2170, 414.3347, 122.1007	Malvidin-3-O-(6-p-coumaroyl) glucoside	Anthocyanin
7.99	C33H40O21	772.70	771.510	230.2559, 459.5033, 313.2480, 122.1008	Kaempferol 3,7,4'-O-triglucoside	Flavonol glycoside
9.41	C30H32O19	696.153	696.439	258.2882, 515.5672, 122.1007	Flavonol base + 3O, O-Hex, O-MalonylHex	Flavonol
*14.17	C33H41O21	773.213	773.547	323.2706, 600.4389, 430.3951, 122.1013	Cyanidin 3,3',5-tri-O-glucoside	Anthocyanin
*14.78	C38H37O18	781.197	780.659	609.2934, 460.3951, 122.1013	( +)-Catechin-Malvidin-3-glucoside	Anthocyanin
*15.11	C45H72O17	885.0452	885.582	593.2973, 690.3885, 429.3881, 122.1011	Kaempferol-rutinosyl-dirhamnoside	Flavonol

*Most abundant compounds identified

Bold numbers indicate the base peaks

Table 2.

Identification and classification of bioactive compounds in extracts of Justicia spicigera, obtained by microwave- assisted extraction (MAE) using chromatography coupled to HPLC–DAD-ESI-QTOF mass spectrometry

TR (min)	Formula	Theoretical mass	[M + H]+ (m/z) cal	ESI–MS/MS Fragments (m/z)	Tentative identification	Type of compound
*2.39	C31H35O19	711.177	711.236	565.1742, 449.1227, 259.1732	Peonidine 3-(6″-malonyl-glucoside) 5-glucoside	Anthocyanin
*2.81	C33H40O18	724.221	725.251	579.1902, 433.1272, 242.1831	Kaempferol-3-rhamnoside-4″-rhamnoside-7-rhamnoside	Flavonoid
7.02	C32H31O14	639.171	639.407	309.2169, 414.3346, 122.1005	Malvidin-3-O-(6-p-coumaroyl) glucoside	Anthocyanin
7.17	C37H39O18	771.214	771.509	291.2054, 448.2836, 122.1005	Peonidine 3-(6″-p-coumarylglucoside)-5-glucoside	Anthocyanin
9.4	C30H32O19	696.153	696.439	258.2882, 515.5672, 122.1007	Flavonol base+3O, O-Hex, O-Malonyl Hex	Flavonol
10.99	C27H31O16	611.529	611.447	295.2375, 395.3035, 122.1007	Cyanidin-3,5-di-O-glucoside	Anthocyanin
11.09	C27H31O16	611.529	611.448	295.2373, 467.2927, 122.1007	Cyanidin-3,5-di-O-glucoside	Anthocyanin
13.55	C38H37O18	781.197	780.657	600.4380, 279.2415, 122.1008	(+)-Catechin-Malvidin-3-glucoside	Anthocyanin
*15.12	C45H72O17	885.0452	885.582	593.2973, 690.3885, 429.3881, 122.1011	Kaempferol-rutinosyl-dirhamnoside	Flavonol

*Most abundant compounds identified

Bold numbers indicate the base peaks

The compounds generally identified in both extraction techniques were flavonoids from the flavonol, flavanol, and anthocyanin families. Furthermore, kaempferol was identified as one of the extracts most abundant compounds using ultrasound and microwave-assisted techniques. Kaempferol 3,7,4′-O-triglucoside has consistently been reported as a major bioactive constituent in aqueous-ethanolic extracts of Justicia spicigera, according to multiple studies.

In the J. spicigera extract obtained via UAE, 11 compounds were tentatively identified, as shown in Table 1. Figure 1 presents the m/z ratios, molecular ions [M + H]⁺, and fragments of some compounds found in J. spicigera extracts, which were considered for the tentative identification of the compounds using HPLC-DAD-ESI-QTOF. Anthocyanins and flavonols, as well as some glycosylated flavonols, were identified. In addition to kaempferol, cyanidin (cyanidin 3,3′,5-tri-O-glucoside) and malvidin (malvidin 3-glucoside-ethyl-catechin), belonging to the anthocyanin group, were among the most abundant compounds.

Fig. 1.

Fragmentation demonstration of some compounds identified by HPLC-DAD-ESI-QTOF. A Epicatechin gallate, B Kaempferol-3-rhamnoside-4″-rhamnoside-7-rhamnoside

On the other hand, J. spicigera extract obtained via MAE, 10 compounds were tentatively identified. The main constituents were glycosylated flavonoids, such as kaempferol, and anthocyanins, including peonidin, malvidin, and cyanidin. Peonidin 3-(6''-malonyl-glucoside) 5-glucoside was the anthocyanin group detected at multiple retention times. This finding aligns with the report by Awad et al. (2015), who indicated that in J. spicigera extracts, peonidin 3,5-diglucoside was the most concentrated anthocyanin. Similarly, the presence of kaempferol derivatives, such as kaempferol-3-rhamnoside-4″-rhamnoside-7-rhamnoside, kaempferol-rutinosyl-dirhamnoside, and kaempferol 3-gentiobioside 7-rhamnoside, is associated with one of the main active compounds found in J. spicigera (Real-Sandoval et al., 2020).

The differences observed in each extraction technique were primarily related to the types of extracted compounds and their respective abundances. These differences may be associated with the compounds' diffusivity from the plant material during both the pretreatment stage and the application of frequency-based (UAE) or wave-based (MAE) extraction techniques. In this way, depending on the extraction technology used, compounds with different chemical structures are obtained. The selectivity of extraction technology could be beneficial in the development of active and intelligent packaging.

Stability of the extracts

Effect of pH and UV–Vis spectroscopy

The UV–visible absorption spectra of the extract obtained via UAE of J. spicigera are shown in Fig. 2A. Under pH conditions ranging from 2 to 5, maximum absorption bands (λmax) were observed in the 306–332 nm range, corresponding to the family of phenolic compounds (Fedenko et al., 2017). At pH levels between 6 and 12, a bathochromic shift in maximum absorption was detected at 315 and 388 nm at pH 12, accompanied by the appearance of new bands at 585 and 673 nm. These bands are characteristic of anthocyanins and may indicate their stability within the pH 6–12 range, while the 673 nm band is associated with chlorophyll-derived pigments (Fedenko et al., 2017). The color of aglycones varies depending on the number of hydroxyl groups, which produce an intense blue hue, as observed in our results within the pH 6–12 range. This effect is intensified as anthocyanidins undergo glycosylation with one or more sugars (Chen and Inbaraj, 2019). These findings correlate with the chemical structure of certain compounds identified via HPLC ESI-QTOF (Table 1), such as cyanidin 3,3′,5-tri-O-glucoside, and malvidin 3-glucoside-ethyl-catechin, which are likely responsible for the coloration observed in the UAE-extracted J. spicigera extract.

Fig. 2.

A UV–Vis absorption spectra of J. spicigera extract, by UAE, at pH value from 2 to 12, B UV–Vis absorption spectra of J. spicigera extract, by MAE, at pH value from 2 to 12

In the case of the extract obtained via MAE, as shown in Fig. 2B, maximum absorption bands (λmax) were observed between 306 and 333 nm (yellow-orange color) at pH 2 and 3. Anthocyanins exhibit a flavylium cation structure under highly acidic conditions, giving them a red–orange hue, as shown in Fig. 3B. Generally, the cationic form hydrates as pH increases to form a colorless carbinol pseudo base with a hemiacetal structure due to a nucleophilic attack in an aqueous medium. This process subsequently leads to the formation of a colorless chalcone, the most unstable form of anthocyanins, ultimately resulting in the loss of pigmentation (Chen & Inbaraj, 2019). However, anthocyanins can interact with other compounds and even with themselves, influencing both their color and structural equilibrium (Enaru et al., 2021).

Fig. 3.

Effect of pH on color parameters in J. spicigera extract; A Luminosity, B Hue angle (°) and C ∆E

In this study, at pH levels above 4, a shift in maximum absorption wavelengths to 314–388 nm was recorded, accompanied by the appearance of an absorption band at 589 nm. The increase in pH could be associated with the observed bathochromic shifts, which may be attributed to the deprotonation of the flavylium cation, a process leading to the formation of a neutral quinoidal base (purple color) under slightly acidic to neutral conditions. The pH can affect the structure of various bioactive compounds, especially flavonoids, because these compounds contain phenolic -OH groups. As pH increases, the compounds are susceptible to deprotonation, producing the purple-blue quinoidal anion, which forms a violet color when interacting with other phenolics (Eze et al., 2022). A study on different flavonoids (quercetin, myricetin, and kaempferol) demonstrated that compound stability under different pH ranges is determined by molecular structure. In this regard, kaempferol was found to be the most stable due to the presence of only one −OH group, whereas myricetin was the least stable, likely due to the presence of three −OH groups (Álvarez-Diduk et al., 2013).

The higher stability observed in the MAE extract could be attributed to acylation with phenolic acids, which enhances anthocyanin stability through steric hindrance (Stintzing and Carle, 2004). This analysis is consistent with the results obtained in the present study. The MAE extract of J. spicigera shows that peonidin 3-(6'-malonylglucoside)-5-glucoside, one of the compounds identified via HPLC ESI-QTOF (Table 2), exhibits greater stability, as evidenced by its blue coloration under acidic pH (4–5) and higher absorbance levels under pH < 4 conditions, likely due to the presence of carboxylic acids in its structure.

Nondestructive methods, such as quantitative color analysis, can be employed to evaluate changes in the colorimetric properties of pigmented extracts exposed to different pH conditions. Changes in the colorimetric parameters of J. spicigera extracts under various pH conditions could be used as potential natural pH indicators.

As shown in Fig. 3A, lightness decreased at pH 6 for the UAE treatment, whereas this decrease occurred at pH 4 for MAE. This behavior corresponds with the observed color shift from red–orange to blue-purple. These color differences are attributed to the predominant compounds recovered by each extraction technology. In both J. spicigera extractions, anthocyanins, particularly peonidin (peonidin 3-(6''-p-coumaroylglucoside)-5-glucoside) and malvidin (malvidin-3-O-(6-p-coumaroyl) glucoside, ( +)-catechin-malvidin-3-glucoside, and malvidin 3-glucoside-ethyl-catechin), were likely responsible for pigmentation and co-pigmentation. The chemical structures of these anthocyanins generate hues ranging from red–orange under acidic conditions to violet at higher pH levels; however, this behavior may vary depending on the anthocyanin’s chemical structure (Enaru et al., 2021).

Figure 3B presents hue angle (°Hue) values as a function of pH. UAE extracts at pH 2–5 exhibited an average hue angle of 47.92°Hue, corresponding to an orange color. At pH ≥ 6, a color shift to pink-purple was observed, with a hue angle of 10.61°Hue. This color change is consistent with the stability of glycosylated anthocyanins identified in the compounds analysis, which were among the predominant components in the UAE extraction. Similarly, the average hue angle for MAE extracts was 49.82°Hue at pH 2 and 3, indicating a red–orange color. However, at pH ≥ 4, the hue angle decreased to an average of 9.64°Hue, indicating a shift toward a blue-purple hue. Similar colorimetric results were reported by Jiménez-González et al. (2023) in J. spicigera extracts, where red–orange tones were observed in acidic media, transitioning to purple-blue in alkaline conditions.

In ultrasound-extracted samples, the highest color changes occurred at pH 6 (ΔE = 14.32), whereas in microwave-extracted samples, the maximum color change occurred at pH 4, with an average ΔE of 14.33. The observed colorimetric differences between extraction techniques highlight the biological functions of chromophores, such as anthocyanins and chlorophylls. Consequently, due to the ability to change color with pH, J. spicigera extract is a valuable source as a freshness indicator in food preservation and as a UV-blocking agent.

Phytochemical composition

Total phenolic compounds (TPC)

Table 3 presents the results of the total phenolic compounds in J. spicigera extracts obtained through UAE and MAE, showing significant differences between treatments (p < 0.05). The J. spicigera extract obtained via MAE exhibited the highest TPC content, with a value of 48.93 ± 2.86 mg GAE/g. In contrast, ultrasound-assisted extraction resulted in a lower TPC yield, reaching 37.40 ± 2.04 mg GAE/g. These differences are attributed to each extraction technology's distinct mechanisms of action, which lead to variations in the recovery and abundance of various bioactive compounds, as demonstrated in the identification of compounds section (HPLC ESI-QTOF).

Table 3.

Values of total phenolic compounds (TPC), total flavonoid compounds (TFC), DPPH scavenging activity, ABTS activity and FRAP activity obtained by ultrasound (UAE) and microwave (MAE) assisted extraction

Extraction method	Total phenolic compounds (mg of GAE/g of dry extract)	Total flavonoid compounds (mg of catechin/g of dry extract)	DPPH (μg of dry extract/mL)	ABTS (µM Trolox/mg of dry extract)	FRAP (µM Trolox/mg of dry extract)
UAE: ultrasound assisted extraction	37.40 ± 2.04a	22.20 ± 0.68a	110.18 ± 3.74a	0.43 ± 0.01a	0.18 ± 0.01a
MAE: microwave assisted extraction	48.93 ± 2.86b	23.46 ± 0.36b	124.47 ± 2.03b	0.45 ± 0.05a	0.21 ± 0.01a

GAE: gallic acid. The values indicated are the mean ± standard deviation. Means with similar letters do not vary significantly, while means with different letters vary according to Tukey’s test (p < 0.05)

The principle of MAE relies on the generation of electromagnetic waves, which induce the movement and friction of solvent molecules. This process increases the temperature and causes physical modification of the matrix (disrupt cell walls and lipoprotein membranes), leading to structural changes that facilitate the release of compounds from the cell membrane (Walayat et al., 2024).

In UAE, the cavitation effect in the liquid medium can generate extremely high temperatures and pressures, which promote the decomposition of solvents and solutes, as well as the depolymerization and degradation of certain compounds, including phenolic compounds (Sayadi et al., 2025).

Furthermore, the TPC values obtained in this study for both extraction methods were significantly higher than those reported by various authors. Awad et al. (2015), using conventional maceration extraction with 95% ethanol, obtained 26.54 mg GAE/g of J. spicigera extract. Similarly, Castro-Alatorre et al. (2021), employing a combination of agitation and an ethanol solvent (40:60% v/v) at 35 °C for 1 h, reported a recovery of 0.33 ± 0.01 mg GAE/g. In summary, microwave-assisted extraction is a more efficient method for recovering phenolic compounds from J. spicigera than ultrasound-assisted extraction.

Total flavonoid compounds (TFC)

Table 3 presents the total flavonoid compounds in J. spicigera extracts obtained through UAE and MAE. The highest flavonoid concentration was achieved using MAE, with a value of 23.46 ± 0.36 mg catechin/g of dry extract, showing significant differences between treatments (p < 0.05). In comparison, UAE resulted in a total flavonoid content of 22.20 ± 0.68 mg catechin/g. These differences highlight the importance of the extraction technique, operating conditions, solvent quantity, and solvent ratio. This mixture promotes cell wall swelling, facilitating the diffusion of target compounds into the solvent. Three types of diffusion processes may occur: (i) Type 1 diffusion: migration of plant components toward the external layer of the tissue. (ii) Type 2 diffusion: migration of components from the tissue into the solvent. (iii) Type 3 diffusion: migration from the external layer into the solvent. (iv) Type 4 diffusion: washing of the cells in the solvent. The resulting solution is then ready for further extraction, such as ultrasound or microwave treatment (Shen et al., 2023).

In MAE, it is crucial to consider the dielectric properties of the mass/solvent ratio to ensure adequate heating of the plant material and the release of target compounds. Understanding these dielectric properties is essential to prevent excessive heating, which could lead to the loss of volatile compounds and degradation of bioactive compounds. In UAE, it is important to note that using an ethanol–water solvent mixture significantly reduces the formation of highly oxidizing compounds. This phenomenon occurs because cavitation induces direct reactions between ethanol and a proportion of water with the plant material. It is worth noting that ethanol is more stable than water in terms of homolytic cleavage (Vinatoru et al., 2017). Our results align with those reported by Castro-Alatorre et al. (2021), using 70% ethanol combined with maceration, reported a flavonoid content of 0.28 ± 0.01 mg catechin/g in J. spicigera extracts. The findings in our study highlight the crucial role of ethanol concentration in recovering total flavonoids from J. spicigera extracts, particularly when combined with efficient extraction techniques such as microwave-assisted extraction.

Antioxidant activity

Free radical scavenging analysis via DPPH

Tests for determining antioxidant activity can be classified according to different chemical reaction mechanisms, including hydrogen atom transfer (HAT) and single electron transfer (SET) (Platzer et al., 2022). A more common way of classification is based on the different methods to assess antioxidant activity: those based on the scavenging activity of a stable free radical (2,2-diphenyl-1-picrylhydrazyl (DPPH) and 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS)) and the reduction of metal ions, such as ferric ion reducing antioxidant potential (FRAP) (Rumpf et al., 2023). Hence, the importance of the evaluation of antioxidant activity by different tests.

Table 3 presents the results of the antioxidant activity of J. spicigera extracts obtained through the DPPH assay. The results show IC₅₀ values with significant differences between treatments (p < 0.05), 110.18 ± 3.74 μg/mL for UAE extraction and 124.47 ± 2.03 μg/mL for MAE. These differences can be attributed to the types of compounds recovered by each extraction technology, as the DPPH method is particularly effective for quantifying lipophilic compounds (Baqueiro-Peña & Guerrero-Beltrán, 2017). Due to the chemical structure of the compounds identified in the UAE extract, such as kaempferol-rutinoside-dirhamnoside, kaempferol 3,7,4′-O-triglucoside, epicatechin gallate, and malvidin 3-glucoside-ethylcatechin, these compounds exhibit lower hydrophilic properties due to their benzene ring structures. Additionally, the antioxidant activity of flavonols and flavones is linked to hydroxyl groups at the 3′ and 4′ positions of ring B and OH at C-3, which allow stable and efficient structures for capturing free radicals (García-Mateos et al., 2012).

On the other hand, the nature of the compounds identified in the MAE extracts, such as kaempferol 3-gentiobioside 7-rhamnoside, peonidin 3-(6″-malonyl-glucoside) 5-glucoside, and malvidin-3-O-(6-p-coumaroyl) glucoside, are more hydrophilic due to their glucosylated structures (Cyboran-Mikołajczyk et al., 2018). Moreover, some researchers indicate that glucosylated compounds are less effective against DPPH. Xiao (2017) demonstrated that 3-O-glucosides of kaempferol exhibited weaker free radical scavenging potential than kaempferol itself. The main difference lies in the abundance with which each extraction technology recovered these compounds. Our results suggest that the recovery of compounds extracted by UAE from J. spicigera could be considered a good source of DPPH radical scavengers compared to MAE.

Determination of antioxidant activity via ABTS

Although the mechanism of the ABTS method is similar to that of the DPPH method, the former is more sensitive, as it can detect both highly active and less active antioxidants. The ABTS method can be used at different pH levels and is soluble in both aqueous and organic solvents, which makes it a suitable method for determining the hydrophilic and lipophilic antioxidant capacity of biological extracts and fluids (Gaber et al., 2023). For this reason, it has sometimes been referred to as “total antioxidant capacity” (Rumpf et al., 2023).

The values obtained from the ABTS^•+ radical inhibition showed Trolox contents with no significant differences between treatments (p > 0.05). With 15 min of MAE, values similar to those obtained with 30 min of UAE were reached (Table 3), with 0.45 ± 0.05 and 0.43 ± 0.01 µM Trolox/mg of dry extract, respectively. The main difference between treatments lies in the extraction time used for each technology. The results obtained in this study were higher than those achieved using conventional technologies, where 0.01863 µM Trolox/mg of dry J. spicigera extract was obtained (Castro-Alatorre et al., 2021). Consequently, implementing new extraction technologies, such as ultrasound and microwaves is keen for the recovery of antioxidant compounds, where treatment time is crucial.

Analysis of iron reducing capacity (FRAP)

Unlike the DPPH and ABTS methods, which are based on the reduction or inhibition of free radicals, the FRAP method is based on the reduction of ferric iron (Fe⁺³) in the FRAP reagent to ferrous iron (Fe⁺²) by the presence of antioxidant compounds (Rumpf et al., 2023).

The results obtained for antioxidant capacity using the FRAP method, as with ABTS, showed no significant differences between the treatments (p > 0.05). The values for antioxidant compounds were 0.18 ± 0.01 and 0.21 ± 0.01 µM Trolox/mg of dry extract for UAE and MAE, respectively (Table 3).

The J. spicigera extract obtained by UAE underwent a 30 min operation at a frequency of 45 kHz. Sonication in the range of ∼20 kHz–2 MHz is classified as low frequency, enhancing cavitation's physical impacts. On the other hand, frequencies higher than 2 MHz favor the chemical impacts of cavitation, considering particle size and the nature of the plant material. During “physical” cavitation, gas and vapor bubbles are formed that generate “hot spots,” a process known as adiabatic compression. Cavitation can be stable or transient, depending on time and conditions. In stable cavitation, bubbles have a prolonged lifespan; in transient cavitation, more bubbles are generated quickly. Moreover, the recovery of bioactive compounds by UAE can be affected or benefited by several factors, such as extraction time, temperature, frequency, and the solid–liquid ratio (plant material/solvent). Two phases distinguish the extraction process: (i) Washing Phase: This phase occurs between 10 and 20 min and represents the interaction time between the plant material and the solvent. During this time, recovery of most of the bioactive compound occurs. (ii) Slow Extraction Phase: This phase involves cavitation and diffusion of compounds by UAE. This process generally lasts between 30 and 100 min (Sabaruddin et al., 2023).

Microwave extraction technology uses dielectric heating for chemical synthesis, significantly reducing extraction times. Other factors affecting this technique include electric fields' polarization and reorientation effects. Dielectric properties directly influence the electric field; solvents must have high dielectric permittivity to absorb the energy generated by microwaves and release the compounds of interest through diffusion. In the case of ethanol, with a dielectric constant of 24.3, uniform heating of the plant material was allowed in this study, achieving shorter extraction times than ultrasound technology (Leonelli and Mason, 2010). The FRAP technique demonstrated that through MAE it is possible to reduce the extraction time by 50% compared to UAE for the recovery of compounds with antioxidant capacity that work through the electron donation mechanism.

The extraction technology is a key factor in the recovery of bioactive compounds from Justicia spicigera, as the mechanism of action of each method influences the type, structure, and abundance of the metabolites obtained. In this study, both ultrasound-assisted extraction (UAE) and microwave-assisted extraction (MAE) techniques enabled the recovery of phenolic compounds, with kaempferol derivatives being the common metabolites identified in both extracts. However, relevant differences were observed in the phytochemical profile: the extract obtained by UAE showed a higher abundance of malvidin and cyanidin derivatives, while peonidin derivatives were predominant in the extract obtained by MAE. Phytochemical analysis revealed a higher content of total phenolic compounds and flavonoids in the extract obtained by MAE. Nevertheless, no statistically significant differences were detected in antioxidant capacity, as evaluated by the ABTS and FRAP methods.

Furthermore, the behavior of the identified compounds, correlated with their stability at different pH ranges, is crucial for their application in food packaging. These findings enable us to conclude that understanding the mechanism of action of extraction technologies, in conjunction with solvent choice, is decisive for guiding the selective recovery of bioactive compounds. Consequently, the obtained extracts show promising properties for application in functional foods, pharmaceutical products, and active packaging systems. Finally, it is suggested that future studies be conducted to evaluate different levels of process parameters (such as time, power, and temperature, among others) for each extraction technique, thereby optimizing efficiency and selectivity in obtaining compounds of interest.

Funding

This work was funded by Secretaría de Investigación y Posgrado, Instituto Politécnico Nacional, 20250602, Erich von Borries Medrano.

Declarations

Conflict of interest

The authors declare no conflict of interest.