Next Article in Journal
Parameter Identification of Five-Phase Squirrel Cage Induction Motor Based on Extended Kalman Filter
Next Article in Special Issue
Optimization of Spray-Drying Conditions of Microencapsulated Habanero Pepper (Capsicum chinense Jacq.) Extracts and Physicochemical Characterization of the Microcapsules
Previous Article in Journal
Multi-Objective Optimization of a Crude Oil Hydrotreating Process with a Crude Distillation Unit Based on Bootstrap Aggregated Neural Network Models
Previous Article in Special Issue
Culturing Important Plants for Sweet Secondary Products under Consideration of Environmentally Friendly Aspects
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Processes
Volume 10
Issue 8
10.3390/pr10081439
processes-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Effect of Plant Growth Regulators on Different Explants of Artemisia ludoviciana under Photoperiod and Darkness Conditions and Their Influence on Achillin Production

by
Mariana Sánchez-Ramos
1,
Samantha Berman-Bahena
1,
Laura Alvarez
2,
Jessica Nayelli Sánchez-Carranza
3,
Antonio Bernabé-Antonio
4,
Angélica Román-Guerrero
1,
Silvia Marquina-Bahena
2,* and
Francisco Cruz-Sosa
1,*
1
Department of Biotechnology Autonomous Metropolitan University-Iztapalapa Campus, Av. Ferrocarril de San Rafael Atlixco 186, Col. Leyes de Reforma 1a. Sección, Alcaldía Iztapalapa, México City 09310, Mexico
2
Chemical Research Center-IICBA, Autonomous University of the State of Morelos, Av. Universidad 1001, Col. Chamilpa, Cuernavaca 62209, Mexico
3
Faculty of Pharmacy, Autonomous University of the State of Morelos, Av. Universidad 1001, Col. Chamilpa, Cuernavaca 62209, Mexico
4
Department of Wood, Pulp and Paper, University Center of Exact Sciences and Engineering, University of Guadalajara, Km 15.5 Guadalajara-Nogales, Col. Las Agujas, Zapopan 45100, Mexico
*
Authors to whom correspondence should be addressed.
Processes 2022, 10(8), 1439; https://doi.org/10.3390/pr10081439
Submission received: 24 June 2022 / Revised: 19 July 2022 / Accepted: 21 July 2022 / Published: 23 July 2022
(This article belongs to the Special Issue Significance and Applicability of Bioactive Compounds from Plants for Innovative Technologies and Processes)
Browse Figures
Review Reports Versions Notes

Abstract

:
Species of the genus Artemisia mainly biosynthesize sesquiterpene lactones. Achillin is a guaianolide-type sesquiterpene lactone isolated from Artemisia ludoviciana; it has shown antibacterial and anti-inflammatory activities. In addition, achillin exhibits a significant chemosensitizing effect on hepatocellular carcinoma cells resistant to paclitaxel (PTX). The objective of this study was to establish a callus culture from different explants under conditions of light and total darkness to produce achillin. To obtain in vitro cultures, explants of leaves, nodes, internodes, and roots were used, and they were cultured in MS medium with 0.1 mg/L of kinetin (KIN) or benzyl amino purine (BAP) and/or naphthaleneacetic acid (NAA), 2,4-dichlorophenoxyacetic acid (2,4-D), indole-3-acetic acid (IAA) and 4-amino-3,5,6-trichloro-2-pyridine carboxylic acid (PIC) at 0.1 and 1.0 mg/L. Of all treatments, internodes with BAP (0.1 mg/L) and PIC (1.0 mg/L) grown under photoperiod showed the best friable callus induction, however, GC-MS analysis showed higher achillin content (1703.05 µg/mL) in leaf calluses with PIC (1.0) and KIN (0.1) under photoperiod, and in node plantlets (1880.01 µg/mL) with PIC (0.1) and BAP (0.1). From 12.34 g of dry leaves of Artemisia ludoviciana, 257 mg of achillin were isolated and purified, which was used as a reference in the quantification of achillin in the in vitro culture.

1. Introduction

The Artemisia genus is a plant that stands out for producing sesquiterpene lactones, and over 500 lactones have been reported. Among the main activities are antitumor, anti-inflammatory, analgesic, antibacterial, antifungal, antiviral and antiparasitic ones [1,2,3,4,5,6]. Artemisia ludoviciana is a plant species popularly known as “estafiate” and it is widely used in traditional Mexican medicine to treat intestinal infections, pain, and inflammation [7]. Some essential oils, flavonoids and different sesquiterpene lactones have been reported for A. ludoviciana [8,9,10,11,12,13,14]. Recent studies report that the compound identified as achillin is one of the leading secondary metabolites for this species [15,16,17]. This compound has a significant effect as a chemosensitizer; when used in combination with an antineoplastic agent, the effect of this agent is enhanced in inducing apoptosis [18]. However, the production of achillin may be limited if it is obtained from wild plants; it is common that the production of secondary metabolites in plants grown in their natural habitat is associated with the stimuli they receive, such as water stress, defense against attack by microorganisms and insects, climatic and season changes, invasion of other species, area where they grow, as well as their growth stages, among other factors [19,20,21,22,23,24,25], and in the case of the achillin metabolite, it is also a compound of a secondary metabolism, which implies that its production depends on the same factors known for this type of compound. Numerous investigations have reported that the yields of secondary metabolites oscillate below 1% in dry weight [26,27,28,29], and given the biological relevance of bioactive compounds [30,31,32], it is feasible to develop strategies that allow their production without compromising the survival of the species [33,34,35]. In this regard, in vitro culture protocols such as callus and suspension cells are of great interest for future applications, which is due to the fact that secondary metabolites of pharmaceutical interest can be produced in a constant and controlled manner, i.e., independently of biotic and abiotic factors. In addition, a higher yield of compounds can be obtained by applying elicitors [36,37,38]. Moreover, this type of culture is a commercially competitive strategy and opens alternatives to the investigation of natural products [39,40,41]. Therefore, this study aimed to establish a callus culture of A. ludoviciana and evaluate the production of achillin and other sesquiterpene lactones.

2. Materials and Methods

2.1. Equipment

Compound achillin was isolated from dry leaves from Artemisia ludoviciana wild plants using column chromatography (silica gel 70–230 mesh; Merck, Darmstadt, Germany). The purity of the compound was observed by UV-Vis spectrophotometers (366 and 254 nm), and after spraying, with lowed by heating. The melting point was obtained on a Thermo Scientific (Waltham, MA, USA) IA9000 series melting point apparatus and was uncorrected. Optical rotation was measured on a 241 digital polarimeter at 25 °C (Perkin Elmer, Waltham, MA, USA) equipped with a sodium lamp (589 nm) and a microcell.
Nuclear magnetic resonance (NMR) of 1H and 13C experiments were recorded on a Bruker AVANCE IIIHD 50 at 500 MHz, using CDCl3 and CD3OD with tetramethylsilane (TMS) as the internal standard. Compound achillin was quantified in in vitro culture using an HP Agilent Technologies 6890 gas chromatograph equipped with an MSD 5973 quadrupole mass detector (HP Agilent, Santa Clara, CA, USA), equipped with a capillary column HP-5MS (length: 30 m; inside diameter: 0.25 mm; film thickness: 0.25 µM). The helium carrier gas was set to the column (1 mL per minute at constant flow). The inlet temperature was set at 250 °C while the oven temperature was initially at 40 °C (held for min) and increased to 280 at 10 °C/min. The mass spectrometer was operated in positive electron impact mode with an ionization energy of 70 eV. Detection was performed in selective ion-monitoring (SIM) mode and peaks were identified and quantitated using target ions (EI-MS m/z (% rel. int.): 246 [M]+ (100), 231 [M-CH3]+, 217 [(29) M-CH2CH3]+, 173 [(44) M-CO2]+, 91 (74).

2.2. Plant Material

Whole wild plants of A. ludoviciana were collected in March 2020 in Cuernavaca, Morelos, México (18°57′29.0″ N, 99°11′57.6″ W). The plant was identified by the Biol. Gabriel Flores Franco and deposited at the Herbarium of the Autonomous University of the State of Morelos (HUMO), Mexico, with the voucher number 33913.

2.2.1. Extraction and Isolation of Achillin from Wild Plant

The leaves of A. ludoviciana were dried in the shade for two days. Then, 12.34 g of dried and ground leaves were subjected to an extraction process by sonication for 30 min using a mixture of dichloromethane and methanol (95:05, v/v). Three cycles of extraction were performed on the same sample. Extracts were combined and concentrated on a rotary evaporator to give a yellow residue (1.4 g). Next, the extract was fractionated according to the described methodology [18]. Finally, 257 mg of the major component was obtained, which was identified as achillin.

2.2.2. Characterization of Achillin Compound

The structure of the isolated compound was confirmed by the analysis of data obtained by Nuclear Magnetic Resonance of Hydrogen and Carbon thirteen, melting point, and optical rotation compared with those already reported for Achillin in the literature.
Achillin was isolated as a crystalline solid: mp 144–145 °C [42]; [α]D: +160 (c 0.5, MeOH) [43]. 1H NMR (500 MHz, CDCl3) δ: 6.04 (s, H–3), 3.70 (t, J = 10.3 Hz, H–6), 3.31 (d, J = 10.2 Hz, H-5), 2.59 (p, J7–11 = 7.7 Hz, H–11), 2.25 (m, H–7), 2.22 (dd, J = 6.3, 1.7 Hz, H–9a), 2.19 (dd, J = 6.3, 1.6 Hz, H–9b), 2.17 (s, CH3–15), 2.08 (s, CH3–14), 1.49 (m, H–8a), 1.39 (m, H–8b), and 1.02 (d, J11–13 = 7.7 Hz, CH3–13). 13C NMR (125 MHz, CDCl3): 196.08 (C–2), 178.64 (C–12), 170.32 (C–4), 152.41 (C–10), 135.55 (C–3), 131.82 (C–1), 83.58 (C–6), 53.00 (C–5), 51.98 (C–7), 39.43 (C–11), 37.67 (C–9), 23.67 (C–8), 21.64 (C–14), 19.88 (C–15) and 10.02 (C–13) [44,45]. Formula: C15H18O3; m/z 246.13. Spectra of 1H and 13C NMR are in Figures S1 and S2. Achillin compound (Figure 1) isolated from the wild plant was used as a reference standard for analyses of in vitro culture extracts.

2.3. Obtaining In Vitro Plantlets of A. ludoviciana

To obtain sufficient aseptic plantlets, they were first multiplied by the in vitro culture using nodal explants from wild plants. Excised nodal explants were washed with faucet water for 2 h and then washed with a commercial detergent solution Axión® for 15 min. Subsequently, segments were superficially disinfected with 70% (v/v) ethanol for 30 s followed by a 5% (v/v) sodium hypochlorite solution (Clorox®, Oakland, CA, USA) for 15 min. Finally, these were rinsed three times with sterile water. Nodal explants were vertically planted into 25×150 mm culture tubes containing 15 mL of MS culture medium [46] without plant growth regulators (PGRs), sucrose (3%, w/v), gelled with Phytagel® (0.25%, w/v) and adjusted to pH 5.7. MS medium was previously autoclaved for 20 min at 121 °C. Cultures were incubated at 25 ± 2 °C under photoperiod conditions for 16 h with white fluorescent light (50-µmol m–2 s–1) for two weeks. Plantlets were micropropagated by subculture every 20 days using the same culture medium.

2.3.1. Evaluation of PGRs in Explants

PGRs were evaluated in different types of explants from 20-day-old in vitro plantlets. Leaf, root, internodes and nodes explants (0.5–1.0 cm) were cultured in MS medium supplemented with auxins and cytokinins as PGRs. Naphthaleneacetic acid (NAA), 2,4-dichlorophenoxyacetic acid (2,4-D), 4-amino-3,5,6-trichloro-2-pyridine carboxylic acid (PIC) or indole-3-acetic acid (IAA) were used as auxins at concentrations of 0, 0.1 and 1.0 mg/L. Each auxin was combined with the cytokinins benzyl amino purine (BAP) or kinetin (KIN) at 0 and 0.1 mg/L. Each treatment consisted of 20 culture tubes per each explant type with one explant per tube, and the experiments were repeated 3 times.

2.3.2. Evaluation of Photoperiod and Total Darkness in Explants

All types of explants were cultured under conditions of photoperiod and total darkness at 25 ± 2 °C. Photoperiod consisted of 16 h with white fluorescent light (50-µmol m–2 s–1) and 8 h of darkness. The other experiment consisted of keeping cultures constantly in total darkness at the same temperature. Data were measured at 20 days of culture and the percentage of morphogenetic response was calculated as the total number of explants from each treatment divided by the total number of explants that had a morphogenetic response and multiplied by 100 [47,48].

2.4. Extraction and Quantification of Achillin from In Vitro Cultures

All 20-day-old treatments showing a morphogenetic response (callus, plantlets, roots or simultaneous induction of calluses and roots) were harvested, dried, ground, and extracted tree times with 100 mL CH2Cl2 each, using an ultrasonic bath for 30 min. The CH2Cl2 extracts were vacuum filtered through Whatman No. 1 filter paper and dried empty. All extracts were monitored by thin layer chromatography using the elution system CH2Cl2:MeOH (95:05) and visualized with a UV-Vis camera. The compound achillin previously isolated from the wild plant was used as a reference standard to calculate the retention factor.
Extracts that revealed evidence of achillin were analyzed by gas chromatography–mass spectrometry (GC-MS). An aliquot of extract (0.5–2.5 mg) was diluted with chloroform (1 mL) and immediately injected into a GC-MS.
For quantitative analysis, a series of solutions containing 2.15, 1.075, 0.5375, 0.2688, 0.1344 and 0.0672 mg/mL of achillin were prepared. Each standard solution was analyzed in triplicate to calculate the ratio of peak area (y) and relative concentration (x); these data were used to construct the linear calibration curve, which showed acceptable linearity with correlation coefficient r2 = 0.9958 (Supplementary Material Figure S3). The results were obtained by analysis in Microsoft Excel 2010. Identification was based on achillin retention time (21.60 min) and MS data (m/z 246.13). The results are expressed in terms of µg of achillin per g of dry biomass.

2.5. Statistical Analysis

The experiments were established in a complementary randomized design; three replicates were used per treatment with 20 explants for each replicate in three independent replicates. The reported data are the means, and the percentage of callus induction was calculated for 4 weeks. One-way ANOVA was used to test the importance of the effects of plant regulators, explant type and their interactions for callus induction. Factors included 26 combinations of hormones and control, four plant segments (leaf, internode, root and nodal segment) and dark and photoperiod conditions. Data analysis was performed with Graph Prism software. The Tukey test was used for mean comparisons (p < 0.05).

3. Results and Discussion

The combinations of cytokines and auxins showed significant differences (p < 0.05) in different explants, whose morphogenetic responses were calluses, roots, seedlings and the formation of callus and roots (callus–roots) that are summarized in Table 1, Table 2 and Table 3. PGR-free explants had no morphogenetic response, turned brown, and died after 5 days of incubation.

3.1. Effect of PGRs in Leaf Explants

The morphogenetic responses for leaf segments were calluses or simultaneous formation of root and callus. Callus was promoted mainly by combining KIN (0.1 mg/L) plus PIC (1.0 mg/L) (Figure 2a), BAP (0.1 mg/L) plus NAA (1.0 mg/L) or BAP (1.0 mg/L) plus PIC (1.0 mg/L) (Figure 2b), which showed statistically non-different induction percentages, with values of 90 ± 5.0, 95 ± 5.0 and 85 ± 5.0%, respectively. The appearance of all calluses was slightly friable; likewise, the BAP (0.1 mg/L) plus NAA (0.1 mg/L) combination promoted callogenesis to a lesser extent than the previous combinations (75 ± 5.0%), and the treatments that revealed less callus induction were the KIN combinations (0.1 mg/L) plus 2,4-D (1.0 mg/L), BAP (0.1 mg/L) plus 2,4-D (0.1 mg/L) (Figure 2c); however, the calluses were compacted in appearance. All calluses showed low friability and the roots turned brown after the second subculture. The morphogenetic responses of leaf explants were favored under photoperiod conditions (Table 1). Simultaneous formation of callus and roots was promoted by combining KIN (0.1 mg/L) plus NAA (1.0 mg/L) (Figure 2d), KIN (0.1 mg/L) plus 2,4-D (0.1 mg/L), KIN (0.1 mg/L) plus 2,4-D (1.0 mg/L) (Figure 2e) or BAP (0.1 mg/L) plus NAA (1.0 mg/L) (Figure 2f), which showed percentages of 73.33 ± 5.77%, 98.33 ± 2.89% and 100%, respectively (Table 2). Other species have reported similar results with the PGRs that were identified in our study, for example, the calluses of the Azadirachta indica species induced in leaf explants were stimulated by the combination KIN and PIC whose relevance lay in the production of the compound azidarichtin with insecticide effect [49]; in the same way, leaf calluses with an antioxidant effect of Eysenhardtia polystachya were induced by the KIN and PIC combination with high cell density and easy disintegration in liquid medium [50]; on the other hand, the combination of BAP and NAA promoted callogenesis in leaf explants of the Artemisia amygdalina species that revealed outstanding antioxidant activity [51]. On another hand, the combination KIN plus 2,4-D was used in the induction of leaf calluses in the species Justicia gendarussa whose calluses exhibited friability and beige coloration [52].

3.2. Effect of PGRs in Nodal Explants

Regarding the nodal explants, they induced formation of calluses or plantlets; ten treatments induced calluses, 6 in photoperiod and 4 in darkness (Table 1 and Table 2), thirteen promoted seedling regeneration, and morphogenetic responses were also favored in the photoperiod. The highest callus induction was generated by the combination KIN (0.1 mg/L) plus 2,4-D (0.1 mg/L) or BAP (0.1 mg/L) plus PIC (1.0 mg/L), achieving percentages of 71 ± 7.64% and 83.33 ± 7.64%; however, the appearance of the callus was friable and slightly brown (Figure 3a). Likewise, combinations of KIN (0.1 mg/L with NAA (1.0 mg/L) promoted 46.67± 2.89% of induction, when combining KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), it achieved 45 ± 5.0% induction (Figure 3b), BAP (0.1 mg/L) with 2,4-D (1.0 mg/L), 31.67 ± 2.89%, and BAP (0.1 mg/L) with PIC (0.1 mg/L), 40 ± 10.00% (Figure 3c); moreover, all calluses were friable. Plantlets formation was the second most induced morphogenetic response of all treatments, and this occurred on nodal explants, mainly combining KIN (0.1 mg/L) plus PIC (1.0 mg/L) with 95 ± 5.00%, BAP (0.1 mg/L) with 2,4-D (0.1 mg/L), achieving an induction of 96.67 ± 2.89%, BAP (0.1 mg/L) with PIC (1.0 mg/L) and BAP (0.1 mg/L) plus IAA (1.0 mg/L), inducing 98.33 ± 2.89% and 91.67 ± 2.89%, respectively (Figure 3d–f). The mentioned combinations were the most outstanding regarding plantlets induction; however, other combinations provided induction, but in a lower percentage (Table 3). Calluses derived from Telfairia occidentalis nodal explants were induced using the combination of BAP plus 2,4-D whose appearance was friable and with low oxidation [53]. The Lycium barbarum callus reported similar results when using the same combination of PGRs and additionally revealed somatic embryogenesis [54]. On the other hand, the calluses induced in the species Cardiospermum halicacabum were promoted by auxin 2,4-D with a smooth and green appearance and also generated as a second morphogenetic response to plant regeneration [55], which could suggest that auxin 2,4-D stimulates callus from nodal explants. The combination of PGRs KIN and NAA stimulated the friable callus induction in Solanum tuberosum, Rauwolfia serpentina, Tinospora formanii [56,57,58].

3.3. Effect of PGRs in Internodal Explants

Internodes induced callus and plantlets formation as morphogenetic responses; 6 treatments were identified that induced calluses in the photoperiod and 5 in darkness (Table 1), while 5 promoted seedling regeneration (Table 3). The combination BAP (0.1 mg/L) with NAA (1.0 mg/L) promoted 100% callus induction (Figure 4a), followed by the combination KIN (0.1 mg/L) with NAA (1.0 mg/L), achieving an induction of 80 ± 10.00%; likewise, combining KIN (0.1 mg/L) plus 2,4-D (1.0 mg/L) had 80% ± 10.00 of induction, and this callus showed a friable appearance (Figure 4b).
Regarding seedling regeneration, the photoperiod conditions were the most appropriate in the morphogenetic response (Table 3). In particular, the combination of KIN and PIC (0.1 mg/L) showed 91.67 ± 2.89% induction, while combining BAP (0.1 mg/L) with IAA (0.1 mg/L), 53.33 ± 2.89% was obtained (Figure 4c). In contrast, the combination of BAP (0.1 mg/L) with IAA (0.1 mg/L) was the only one that produced seedlings under dark conditions (Figure 4d). These results suggest that the cytokinin BAP stimulates the morphogenetic response. The induction response under dark conditions was similar to those obtained with the photoperiod mg/L (Table 1). The combination of KIN (0.1 mg/L) and PIC (0.1 mg/L) exhibited 91.67% ± 2.89 of plantlets induction, while for BAP (0.1 mg/L) with IAA (0.1 mg/L), only 53.33% ± 2.89 was obtained (Figure 4c). Other combinations of BAP with PIC or IAA also induced plantlets (Figure 4d), suggesting that BAP has an enhanced induction compared to cytokinin KIN; furthermore, under dark conditions, morphogenetic responses were significantly inhibited (Table 3).
The internodal explants of Asteracantha longifolia, Thymus pesicus, Dianthus caryophyllus were induced to callus by PGRs NAA and BAP with a friable appearance [59,60,61]; on the other hand, the combination of KIN and NAA promoted friable calluses in internodal explants of Solanum tuberosum [62], PGRs 2,4-D and BAP promoted a smooth-appearing callus in internodal explants of Capsicum annuum [63]. Additionally, the combination of BAP with IAA stimulated the regeneration of seedlings in internodal explants of Solanum nigrum [64], while the internodal explants of Solanum trilobatum L. induced seedling regeneration when using BAP and IAA [65].

3.4. Effect of PGRs in Root Explants

As seen in Table 1, only 7 treatments induced callus induction (four under photoperiod conditions and three in darkness). Root explants showed 100% of callus induction when KIN (0.1 mg/L) with 2,4-D (1.0 mg/L) was added to culture medium (Figure 5a), and these same PGRs induced friable calluses in root explants of Withania sommifera, Cuminum cyminum and Oryza sativa species [66,67,68]. Other treatments such as KIN (0.1 mg/L) with NAA (1.0 mg/L), KIN (0.1 mg/L) with PIC (1.0 mg/L) and BAP (0.1 mg/L) with PIC (1.0 mg/L) exhibited 80% ± 5.00 of induction (Figure 5b), whose appearance was friable during the first two subcultures in periods of 25 days; rather, they turned brown and compact in appearance; similarly, the calluses of root explants of Lathyrus sativus when the PGRs NAA and KIN [69]. The only treatment that induced simultaneous formation of callus and root (Table 2) occurred in the treatment with KIN (0.1 mg/L) plus 2,4-D (0.1 mg/L) under dark conditions, but they were also compact in appearance and brown (Figure 5c).
All the explants favored callus induction due to PGRs; however, the internodal explants showed a greater abundance of calluses compared to other treatments. In addition, the combination of BAP (0.1 mg/L) with PIC (1.0 mg/L) presented better friable callus. The other treatments were also able to form calluses, but these were compact in appearance, brown and of little growth. Plant growth regulators affect growth, differentiation, regeneration and production of secondary metabolites in plants and tissue cultures [70,71,72,73]. Cytokinins mainly stimulate the production of phenolic compounds in Artemisia species. For instance, A. annua showed significant accumulation of the lactone artemisinin in plantlets cultured in MS medium with KIN/NAA [74]. Leaf explants of A. absinthium induced calluses when exposed to KIN/NAA combination, and they also showed shoot regeneration after 60 days of age, although most calluses did not survive the subcultures and their growth was slow; however, BAP/NAA-induced calluses were friable and some even showed organogenesis indirectly [75]. In another study, A. absinthium leaf explants induced calluses using thidiazuron, revealing a high frequency of callogenesis and high accumulation of flavonoids, carotenoids and phenols [76]. On the other hand, calluses of Lippia multiflora were induced with zeatin; however, by combining with PIC, callus induction, friability and proliferation were improved [77].
These results show that for A. ludoviciana, internodal explants are the most suitable for obtaining friable calluses under photoperiod conditions. This result is similar to that reported in A. amygdalina where internodes and leaf explants successfully induced callus formation [51]. Leaf explants of A. absinthium, A. annua and A. dracunculus have been shown in other studies to be the best source of explants for callus induction [75,78,79,80], while for A. scoparia, the petiole explant has been better for callus induction [81], which indicates that in addition to the regulator and the explant, the genotype is an important factor in the formation of calluses. These morphogenetic responses are associated with the applied plant growth regulators.

3.5. Effect of Photoperiod on Morphogenetic Response

The photoperiod was one of the factors that influenced the different desirable morphogenetic responses for this study (Table 1, Table 2 and Table 3). Irradiance has favorable effects on plant cells, tissue growth and the biosynthesis of secondary metabolites [82,83,84,85]. The results of our study are similar to those reported for callus cultures of Justicia gendarussa, where there was a stimulation in the production of terpene compounds when cultivated under photoperiod conditions [83]. Likewise, Gerbera jamesonii callus cultures induced under photoperiod conditions showed greater robustness than those in darkness [86]. On the other hand, Abelmoschus esculentus callus cultures showed a greater accumulation of biomass and secondary compounds under photoperiod conditions [87].

3.6. Achillin Quantification in In Vitro Cultures Extracts

We previously isolated and purified the achillin compound from the dichloromethane: methanol (95:05) leaf extract from A. ludoviciana wild plant. The pure compound (Figure 6a–c) was used as a reference for quantification of achillin in dichloromethane extracts from the in vitro culture. Chromatograms, as well as data about the quantification method, are reported in the Supplementary Material (Figures S1–S3).
Of the 216 treatments evaluated, 42 treatments induced calluses (13 of leaf, 11 of nodal, 11 of internodes and 7 of roots explants) and 17 treatments regenerated plantlets (12 of nodal and 5 of internodal explants). The biomass of the subcultured treatments every 20 days for four months was dried in an oven at 60 °C and extracted with CH2Cl2 by sonication. The chromatographic profile performed on all samples (Figure S3, Supplementary Material) revealed that only 10 treatments showed achillin, and they are shown in Table 4.
Notably, the plantlets revealed a higher content of achillin, while the calluses exhibit a lower proportion; however, it is the first study of the A. ludoviciana species that has reported this outstanding result, which undoubtedly shows that callus cultures are a viable alternative for producing biologically active compounds.
Other species of the same genus have reported the production of compounds of biological interest: by means of callus cultures of the species A. spicigera, volatile organic compounds were produced; likewise, callus cultures of A. scoparia produced antimalarial compounds [84,88].

4. Conclusions

The present study demonstrated the capacity of the callus cultures of A. ludoviciana for the biosynthesis of achillin, a guaianolide-type sesquiterpene lactone with a significant effect as a chemo-sensitizer in cells of hepatocellular carcinoma, which present resistance to paclitaxel. In the protocol established for the callus culture, it was found that the internode explants cultured in the MS medium supplemented with plant growth regulators BAP and PIC exhibited high friability under photoperiod conditions. In addition, GC-MS analysis of the dichloromethane extract showed the presence of the compound aquiline, demonstrating that callus cultures of A. ludoviciana biosynthesize sesquiterpene lactones. These results represent an advantage in the production of bioactive compounds in a sustainable way using a biotechnological crop, and in the future, it could be produced on a larger scale using bioreactors or elicitors.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/pr10081439/s1, Figure S1: Achillin compound (RNM 1H (500 MHz), CDCl3; Figure S2: Achillin compound RNM 13C (125 MHz), CDCl3; Figure S3: Calibration curve using achillin compound isolated from a wild plant.

Author Contributions

Conceptualization, M.S.-R. and F.C.-S.; formal analysis, A.R.-G., A.B.-A. and S.M.-B.; investigation, L.A., S.B.-B. and M.S.-R.; writing—original draft preparation, M.S.-R., A.B.-A. and J.N.S.-C.; visualization, A.R.-G. and L.A.; supervision, F.C.-S. and S.M.-B.; funding acquisition, F.C.-S. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available on request from the corresponding authors.

Acknowledgments

We thank María Gregoria Medina Pintor of the Chemical Research Center (UAEM) for her kind support in providing laboratory assistance and gas chromatograph-mass spectrometry procedures.

Conflicts of Interest

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

References

  1. Ivanescu, B.; Miron, A.; Corciova, A. Sesquiterpene lactones from Artemisia Genus: Biological activities and methods of analysis. J. Anal. Methods Chem. 2015, 2015, 247685. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Merfort, I. Perspectives on sesquiterpene lactones in inflammation and cancer. Curr. Drug Targets 2011, 12, 1560–1573. [Google Scholar] [CrossRef] [PubMed]
  3. Moujir, L.; Callies, O.; Sousa, P.M.C.; Sharopov, F.; Seca, A.M.L. Applications of sesquiterpenes lactones: A review of some potential success cases. Appl. Sci. 2020, 10, 3001. [Google Scholar] [CrossRef]
  4. Palacios-Espinosa, J.F.; Núñez-Aragón, P.N.; Gomez-Chang, E.; Linares, E.; Bye, R.; Romero, I. Anti-helicobacter pylori activity of Artemisia ludoviciana subsp. mexicana and two of its bioactive components, estafiatin and eupatilin. Molecules 2021, 26, 3654. [Google Scholar] [CrossRef]
  5. Abad, M.M.J.; Bedoya, D.O.L.M.; Apaza, T.L.; Bermejo, B.P. The Artemisia L. Genus: A review of bioactive sesquiterpene lactones. Stud. Nat. Prod. Chem. 2012, 37, 43–65. [Google Scholar]
  6. Zhang, S.; Won, Y.-K.; Ong, C.N.; Shen, H.-M. Anti-cancer potential of sesquiterpene lactones: Bioactivity and molecular mechanisms. Curr. Med. Chem. Anticancer Agents 2005, 5, 239–249. [Google Scholar] [CrossRef] [PubMed]
  7. Argueta, A.; Cano, L.M.; Rodarte, M.E. Atlas de las Plantas de la Medicina Tradicional Mexicana, Tomo I–III, 1st ed.; Instituto Nacional Indigenista: Mexico City, Mexico, 1994. [Google Scholar]
  8. Epstein, W.W.; Ubben, J.E.E. Anthemidin, a new sesquiterpene lactone from Artemisia ludoviciana. J. Nat. Prod. 1979, 42, 279–281. [Google Scholar] [CrossRef]
  9. Jakupovic, J.; Tan, R.X.; Bohlmann, F.; Boldt, P.E.; Jia, Z.J. Sesquiterpene lactones from Artemisia ludoviciana. Phytochemistry 1991, 30, 1573–1577. [Google Scholar] [CrossRef]
  10. Ruiz-Cancino, A.; Cano, A.E.; Delgado, G. Sesquiterpene lactones and flavonoids from Artemisia ludoviciana ssp. Mexicana. Phytochemistry 1993, 33, 1113–1115. [Google Scholar] [CrossRef]
  11. Said, F.S.; Ramos, G.M.C.; Mata, C.B.D.; Vargas, V.J.; Villarreal, T.L. In vitro antiprotozoal activity of the leaves of Artemisia ludoviciana. Fitoterapia 2005, 76, 466–468. [Google Scholar] [CrossRef]
  12. Anaya-Eugenio, G.D.; Rivero-Cruz, I.; Rivera-Chávez, J.; Mata, R. Hypoglycemic properties of some preparations and compounds from Artemisia ludoviciana Nutt. J. Ethnopharmacol. 2014, 155, 416–425. [Google Scholar] [CrossRef] [PubMed]
  13. Anaya-Eugenio, G.D.; Rivero-Cruz, I.; Bye, R.; Linares, E.; Mata, R. Antinociceptive activity of the essential oil from Artemisia ludoviciana. J. Ethnopharmacol. 2016, 179, 403–411. [Google Scholar] [CrossRef] [PubMed]
  14. Estrada-Soto, S.; Sánchez-Rencillas, A.; Navarrete-Vázquez, G.; Castillo-España, P.; Villalobos-Molina, R.; Ibarra-Barajas, M. Relaxant effects of Artemisia ludoviciana on isolated rat smooth muscle tissues. J. Ethnopharmacol. 2012, 139, 513–518. [Google Scholar] [CrossRef]
  15. Ohno, N.; Gershenzon, J.; Roane, C.; Mabry, T.J. 11,13-dehydrodesacetylmatricarin and other sesquiterpene lactones from Artemisia ludoviciana Var. Ludoviciana and the identity of artecanin and chryrsartemin B. Phytochemistry 1980, 19, 103–106. [Google Scholar] [CrossRef]
  16. Rivero-Cruz, I.; Anaya-Eugenio, G.; Pérez-Vásquez, A.; Martínez, L.; Mata, R. Quantitative analysis and pharmacological effects of Artemisia ludoviciana aqueous extract and compounds. Nat. Prod. Commun. 2017, 12, 1531–1534. [Google Scholar] [CrossRef] [Green Version]
  17. Romero, J.L.; Sosa, C.M.; Burgoa, G.L.; Leal, A.C.; El Kassis, E.G.; Rodríguez, E.B.; Juárez, G.A.; Hernández, L.R.; Bach, H.; Juárez, Z.N. Antimycobacterial, cytotoxic, and anti-inflammatory activities of Artemisia ludoviciana. J. Ethnopharmacol. 2022, 293, 115249. [Google Scholar] [CrossRef] [PubMed]
  18. Sánchez-Carranza, J.N.; González-Maya, L.; Razo-Hernández, R.S.; Salas-Vidal, E.; Nolasco-Quintana, N.Y.; Clemente-Soto, A.F.; García-Arizmendi, L.; Sánchez-Ramos, M.; Marquina, S.; Alvarez, L. Achillin increases chemosensitivity to paclitaxel, overcoming resistance and enhancing apoptosis in human hepatocellular carcinoma cell line resistant to paclitaxel (Hep3B/PTX). Pharmaceutics 2019, 11, 512. [Google Scholar] [CrossRef] [Green Version]
  19. Ramakrishna, A.; Ravishankar, G.A. Influence of abiotic stress signals on secondary metabolites in plants. Plant Signal. Behav. 2011, 6, 1720–1731. [Google Scholar]
  20. Rhodes, M.J.C. Physiological roles for secondary metabolites in plants: Some progress, many outstanding problems. Plant Mol. Biol. 1994, 24, 1–20. [Google Scholar] [CrossRef]
  21. Vasconsuelo, A.; Boland, R. Molecular aspects of the early stages of elicitation of secondary metabolites in plants. Plant Sci. 2007, 172, 861–875. [Google Scholar] [CrossRef]
  22. Shitan, N. Secondary metabolites in plants: Transport and self-tolerance mechanisms. Biosci. Biotechnol. Biochem. 2016, 80, 1283–1293. [Google Scholar] [CrossRef] [Green Version]
  23. Soni, U.; Brar, S.; Gauttam, V.K. Effect of seasonal variation on secondary metabolites of medicinal plants. Int. J. Pharm. Sci. Res. 2015, 6, 3654–3662. [Google Scholar]
  24. Pant, P.; Pandey, S.; Dall’Acqua, S. The influence of environmental conditions on secondary metabolites in medicinal plants: A literature review. Chem. Biodivers. 2021, 18, e2100345. [Google Scholar] [CrossRef]
  25. Mishra, T. Climate change and production of secondary metabolites in medicinal plants: A review. Int. J. Herb. Med. 2016, 4, 27–30. [Google Scholar]
  26. Li, Y.; Kong, D.; Fu, Y.; Sussman, M.R.; Wu, H. The effect of deveplomental and environmental factors on secondary metabolites in medicinal plants. Plant Physiol. Biochem. 2020, 148, 80–89. [Google Scholar] [CrossRef]
  27. Wawrosch, C.; Zotchev, S.B. Production of bioactive plant secondary metabolites through in vitro technologies-status and outlook. Appl. Microbiol. Biotechnol. 2021, 105, 6649–6668. [Google Scholar] [CrossRef] [PubMed]
  28. Singh, A.; Dwivedi, P. Methyl-jasmonate and salicylic acid as potent elicitors for secondary metabolite production in medicinal plants: A review. J. Pharmacog. Phytochem. 2018, 7, 750–757. [Google Scholar]
  29. Narayani, M.; Srivastava, S. Elicitation: A stimulation of stress in in vitro plant cell/tissue cultures for enhancement of secondary metabolite. Phytochem. Rev. 2017, 16, 1227–1252. [Google Scholar] [CrossRef]
  30. González, I.F.M.; González, D.E.F.; Morena, V.C. Secondary metabolites in plants: Main classes, phytochemical analysis and pharmacological activities. Bionatura 2019, 4, 1000–1009. [Google Scholar] [CrossRef]
  31. Savithramma, N.; Linga, M.R.; Suhrulatha, D. Screening of medicinal plants for secondary metabolites. Middle East J. Sci. Res. 2011, 8, 579–584. [Google Scholar]
  32. Jain, C.; Khatana, S.; Vijayvergia, R. Bioactivity of secondary metabolites of various plants: A review. Int. J. Pharm. Sci. Res. 2018, 10, 494–504. [Google Scholar]
  33. Vanisree, M.; Lee, C.-Y.; Lo, S.-F.; Nalawade, S.M.; Lin, C.Y.; Tsay, H.-S. Studies on the production of some important secondary metabolites from medicinal plants by tissue cultures. Bot. Bull. Acad. Sin. 2004, 45, 1–22. [Google Scholar]
  34. Yue, W.; Ming, Q.-L.; Lin, B.; Rahman, K.; Zheng, C.-J.; Han, T.; Qin, L.-P. Medicinal plant cell suspension cultures: Pharmaceutical applications and high-yielding strategies for the desired secondary metabolites. Crit. Rev. Biotechnol. 2014, 1, 215–232. [Google Scholar] [CrossRef] [PubMed]
  35. Isah, T.; Umar, S.; Mujib, A.; Prasad, M.S.; Rajasekharan, P.E.; Zafar, N.; Frukh, A. Secondary metabolism of pharmaceuticals in the plant in vitro cultures: Strategies, approaches, and limitations to achieving higher yield. Plant Cell Tissue Organ Cult. (PCTOC) 2017, 132, 239–265. [Google Scholar] [CrossRef]
  36. Mulabagal, V.; Tsay, H.S. Plant cell cultures—An alternative and efficient source for the production of biologically important secondary metabolites. Int. J. Appl. Sci. Eng. 2004, 2, 29–48. [Google Scholar]
  37. Smetanska, I. Production of secondary metabolites using plant cell cultures. Adv. Biochem. Eng./Biotechnol. 2008, 111, 187–228. [Google Scholar]
  38. Ahuja, A.; Tripathi, M.K.; Tiwari, S.; Tripathi, N.; Tiwari, G.; Mishra, N.; Bhargav, S.; Tiwari, S. Recent advancements on callus and cell suspension cultures: An effectual reserve for the production of pharmaceutically significant metabolites. Curr. Asp. Pharm. Res. Dev. 2021, 6, 96–111. [Google Scholar]
  39. Chattopadhyay, S.; Farkya, S.; Srivastava, A.K.; Bisaria, V.S. Bioprocess considerations for production of secondary metabolites by plant cell suspension cultures. Biotechnol. Bioprocess. Eng. 2002, 7, 138–149. [Google Scholar] [CrossRef]
  40. Moscatiello, R.; Baldan, B.; Navazio, L. Plant cell suspension cultures. In Plant Mineral Nutrients Methods and Protocols; Maathius, F.J.M., Ed.; Springer Protocols: Berlin/Heidelberg, Germany, 2012; pp. 77–93. [Google Scholar]
  41. Wu, T.; Kerbler, S.M.; Fernie, A.R.; Zhang, Y. Plant cell cultures as heterologous bio-factories for secondary metabolite production. Plant Commun. 2021, 2, 100235. [Google Scholar] [CrossRef]
  42. Marx, J.N.; White, E.H. The stereochemistry and synthesis of Achillin. Tetrahedron 1969, 25, 2117–2120. [Google Scholar] [CrossRef]
  43. Ho, C.; Ju, E.C.; Soo, G.Y.; Kim, K.-M.; Yong, S.I. Desacetylmatricarin, an Anti-allergic component from Taraxacum platycarpum. Planta Med. 1998, 64, 557–578. [Google Scholar] [CrossRef] [PubMed]
  44. Martinez, V.M.; Muñoz-Zamora, A.; Joseph-Nathan, P. Conformational analysis of achillin and leukodin. J. Nat. Prod. 1988, 51, 221–228. [Google Scholar] [CrossRef]
  45. Balboul, B.A.A.A.; Ahmed, A.A.; Otsuka, H.; Bando, M.; Kido, M.; Takeda, Y. A guaianolide and a germacranolide from Achillea santolina. Phytochemistry 1997, 46, 1045–1049. [Google Scholar] [CrossRef]
  46. Murashige, T.; Skoog, F. A revised medium for rapid growth and bio Assays with Tobacco tissue cultures. Physiol. Plant 1962, 15, 473–497. [Google Scholar] [CrossRef]
  47. Tariq, M.; Ali, G.; Hadi, F.; Ahmad, S.; Ali, N.; Ali, S.A. Callus induction and in vitro plant regeneration of Rice (Oryza sativa L.) under various conditions. Pak. J. Biol. Sci. 2008, 11, 255–259. [Google Scholar] [CrossRef] [PubMed]
  48. Islam, M.M.; Ahmed, M.; Mahaldar, D. In vitro callus induction and plant regeneration in seed explants of Rice (Oryza Sativa L.). J. Agric. Biol. Sci. 2005, 1, 72–75. [Google Scholar]
  49. Farjaminezhad, R.; Garoosi, G.-A. New biological trends on cell and callus growth and azadirachtin production in Azadirachta indica. Biotech 2019, 9, 309. [Google Scholar] [CrossRef]
  50. Bernabé-Antonio, A.; Maldonado-Magaña, A.; Ramírez-López, C.B.; Salcedo-Pérez, E.; Meza-Contreras, J.C.; González-García, Y.; López-Dellamary, T.F.A.; Cruz-Sosa, F. Establishment of callus and cell suspension cultures of Eysenhardtia polystachya (Ortega) and fungistatic activity of their extracts. S. Afr. J. Bot. 2017, 112, 40–47. [Google Scholar] [CrossRef]
  51. Rasool, R.; Ganai, B.A.; Kamili, A.N.; Akbar, S. Antioxidant potential in callus culture of Artemisia amygdalina Decne. Nat. Prod. Res. 2012, 26, 2103–2106. [Google Scholar]
  52. Thomas, T.D.; Yoichiro, H. In vitro propagation for the conservation of a rare medicinal plant Justicia gensarussa Burm. f. by nodal explants and shoot regeneration from callus. Acta Physiol. Plant 2010, 32, 943–950. [Google Scholar] [CrossRef]
  53. Sakpere, A.M.A.; Ajayi, S.A.; Adelusi, A.A. Effect of growth regulators and explant types on callus induction in Telfairia occidentalis Hook F. Afr. J. Biotechnol. 2014, 13, 2015–2021. [Google Scholar] [CrossRef] [Green Version]
  54. Osman, N.I.; Awal, A.; Sidik, N.J.; Abdullah, S. Callus induction and somatic embryogenesis from leaf and nodal explants of Lycium barbarum L. (Goji). Biotechnology 2013, 12, 36–45. [Google Scholar] [CrossRef] [Green Version]
  55. Thomas, T.D.; Maseena, E.A. Callus induction and plant regeneration in Cardiospermum halicacabum Linn. an important medicinal plant. Sci. Hortic. 2006, 108, 332–336. [Google Scholar] [CrossRef]
  56. Kumlay, A.M.; Ercisli, S. Callus induction, shoot proliferation and root regeneration of potato (Solanum tuberosum L.) stem node and leaf explants under long-day conditions. Biotechnol. Biotecnol. Equip. 2015, 29, 1075–1084. [Google Scholar] [CrossRef] [Green Version]
  57. Pandey, V.P.; Kudakasseril, J.; Cherian, E.; Patani, G. Comparison of two methods for in vitro propagation of Rauwolfia serpentina from nodal explants. Indian Drugs 2007, 44, 514–519. [Google Scholar] [CrossRef]
  58. Dharmapal, S.P.; Najla, M.; Swetha, E.S.; Udayan, P.S.; Elyas, K.K. Callus induction and organogenesis from Tinospora formanii Udayan and Pradeep: A rare endemic plant. Trop. Plant Res. 2017, 4, 71–76. [Google Scholar] [CrossRef]
  59. Kumar, M.S.; Nandi, S.C. High frequency plant regeneration with histological analysis of organogenic callus from internode explants of Asteracantha longifolia Nees. J. Genet. Eng. Biotechnol. 2015, 13, 31–37. [Google Scholar] [CrossRef] [Green Version]
  60. Bakhtiar, Z.; Mirjalili, M.H.; Sonboli, A. In vitro callus induction and micropropagation of Thymus persicus (Lamiaceae), an endangered medicinal plant. Crop. Breed. App. Biotechnol. 2016, 16, 48–54. [Google Scholar] [CrossRef]
  61. Arif, M.; Rauf, S.; Din, A.U.; Rauf, M.; Afrasiab, H. High frequency plant regeneration from leaf derived callus of Dianthus caryophyllus L. Am. J. Plant Sci. 2014, 5, 2454–2463. [Google Scholar] [CrossRef] [Green Version]
  62. Shirin, F.; Hossain, M.; Kabir, M.F.; Roy, M.; Sarker, S.R. Callus induction and plant regeneration from internodal and leaf explants of four potato (Solanum tuberosum L.) cultivars. World J. Agric. Sci. 2007, 3, 01–06. [Google Scholar]
  63. Khan, H.; Siddique, I.; Anis, M.; Khan, P.R. In vitro organogenesis from internode derived callus cultures of Capsicum annuum L. J. Plant Biochem. Biotechnol. 2011, 20, 84–89. [Google Scholar] [CrossRef]
  64. Sridhar, T.M.; Naidu, C.V. An efficient callus induction and plant regeneration of Solanum nigrum (L.)—An important antiulcer medicinal plant. J. Phytol. 2011, 3, 23–28. [Google Scholar]
  65. Arockiasamy, D.I.; Muthukumar, B.; Natarajan, E.; Britto, S.J. Plant regeneration from node and internode explants of Solanum trilobatum L. Plant Tissue Cult. 2002, 12, 93–97. [Google Scholar]
  66. Rani, G.; Virk, G.S.; Nagpal, A. Callus induction and plantlet regeneration in Withania somnifera (L.) dunal. In Vitro. Cell. Dev. Biol. Plant 2003, 39, 468–474. [Google Scholar] [CrossRef]
  67. Soorni, J.; Kahrizi, D. Effect of genotype, explant type and 2,4-D on cell differentiation and callus induction in Cumin (Cuminum cyminum L.) medicinal plant. J. Appl. Biotechnol. Rep. 2015, 2, 265–270. [Google Scholar]
  68. Bano, S.; Jabeen, M.; Rahim, F.; Ilahi, I. Callus induction and regeneration in seed explants of rice (Oryza sativa CV. Swat-II). Pak. J. Bot. 2005, 37, 829–836. [Google Scholar]
  69. Roy, P.K.; Barat, G.K.; Mehta, S.L. In vitro plant regeneration from callus derived from root explants of Lathyrus sativus. Plant Cell Tissue Organ Cult. 1992, 29, 135–138. [Google Scholar] [CrossRef]
  70. Zhong, J.J.; Bai, Y.; Wang, S.J. Effects of plant growth regulators on cell growth and ginsenoside saponin production by suspension cultures of Panax quinquefolium. J. Biotechnol. 1996, 45, 227–234. [Google Scholar] [CrossRef]
  71. Farhadi, N.; Panahandeh, J.; Azar, A.M.; Salte, S.A. Effects of explant type, growth regulators and light intensity on callus induction and plant regeneration in four ecotypes of Persian shallot (Allium hirtifolium). Sci. Hortic. 2017, 218, 80–86. [Google Scholar] [CrossRef]
  72. Gao, F.; Peng, C.; Wang, H.; Shen, H.; Yang, L. Selection of culture conditions for callus induction and proliferation by somatic embryogenesis of Pinus koraiensis. J. For. Res. 2021, 32, 483–491. [Google Scholar] [CrossRef]
  73. Guo, G.; Ryong, B.J. Explant, medium, and plant growth regulator (PGR) affect induction and proliferation of callus in Abies koreana. Forests 2021, 12, 1388. [Google Scholar] [CrossRef]
  74. Fulzele, D.P.; Sipahimalani, A.T.; Heble, M.R. Tissue cultures of Artemisia annua: Organogenesis and Artemisinin production. Phytother Res. 1991, 5, 149–153. [Google Scholar] [CrossRef]
  75. Nin, S.; Morosi, E.; Schiff, S.; Bennici, A. Callus cultures of Artemisia absinthium L.: Initiation, growth optimization and organogenesis. Plant Cell Tissue Organ Cult. 1996, 45, 67–72. [Google Scholar] [CrossRef]
  76. Tariq, U.; Ali, M.; Abbasi, B.H. Morphogenic and biochemical variations under different spectral lights in callus cultures of Artemisia absinthium L. J. Photochem. Photobiol. B: Biol. 2014, 130, 264–271. [Google Scholar] [CrossRef] [PubMed]
  77. André, S.B.; Mongimaké, K.; Modeste, K.K.; Edmond, K.K.; Tchoa, K.; Hilare, K.T.; Yatty, K.Y. Effects of plant growth regulators and carbohydrates on callus induction and proliferation from leaf explant of Lippia multiflora Moldenke (Verbenacea). Int. J. Agric. Crop Sci. 2015, 8, 118–127. [Google Scholar]
  78. Ali, M.; Haider, B.A. Thidiazuron-induced changes in biomass parameters, total phenolic content, and antioxidant activity in callus cultures of Artemisia absinthium L. Appl. Biochem. Biotechnol. 2014, 172, 2363–2376. [Google Scholar] [CrossRef] [PubMed]
  79. Ibrahim, A.K.; Ahmed, S.A. Efficient callus induction, plant regeneration and estragole estimation in Tarragon (Artemisia dracunculus L.). J. Essent. Oil Res. 2011, 23, 16–20. [Google Scholar] [CrossRef]
  80. Dangash, A.; Ram, M.; Niranjan, R.; Bharillya, A.; Misra, H.; Pandya, N.; Jain, D.C. In vitro selection and hormonal regulation in cell culture of Artemisia annua L. Plant. JSM Cell Dev. Biol. 2015, 3, 1013. [Google Scholar]
  81. Aslam, N.; Zia, M.; Chaudhary, M.F. Callogenesis and direct organogenesis of Artemisia scoparia. Pak. J. Biol. Sci. 2006, 9, 1783–1786. [Google Scholar] [CrossRef]
  82. Ouyang, J.; Wang, X.; Zhao, B.; Wang, Y. Light intensity and spectral quality influencing the callus growth of Cistanche deserticola and biosynthesis of phenylethanoid glycosides. Plant Sci. 2003, 165, 657–661. [Google Scholar] [CrossRef]
  83. Wahyuni, D.K.; Huda, A.; Faizah, S.; Purnobasuki, H.; Wardojo, B.P.E. Effects of light, sucrose concentration and repetitive subculture on callus growth and medically important production in Justicia gendarussa Burm. f. Biotechnol. Rep. 2020, 27, e00473. [Google Scholar] [CrossRef] [PubMed]
  84. Ghorbani, S.; Kosari-Nasab, M.; Mahjouri, S.; Talebpour, A.H.; Movafeghi, A.; Maggi, F. Enhancement of in vitro production of volatile organic compounds by shoot differentiation in Artemisia spicigera. Plants 2021, 10, 208. [Google Scholar] [CrossRef]
  85. Morini, S.; D’Onofrio, C.; Bellocchi, G.; Fisichella, M. Effect of 2,4-D and light quality on callus production and differentiation from in vitro cultured quince leaves. Plant Cell Tissue Organ Cult. 2000, 63, 47–55. [Google Scholar] [CrossRef]
  86. Mohlakola, E.M.; Cheng, C.; Lin, Y.; Guo, R.; Min, K.T.; Chen, Y.; Lai, Z. Effects of 2,4-Dichlorophenoxy acetic acid and light on growth of Gerbera (Gerbera jamesonii cv. Daxueju) callus. ACS Agric. Sci. Technol. 2017, 18, 385–393. [Google Scholar]
  87. Irshad, M.; Debnath, B.; Mitra, S.; Arafat, Y.; Li, M.; Sun, Y.; Qiu, D. Accumulation of anthocyanin in callus cultures of red-pod okra [Abelmoschus esculentus (L.) Hongjiao] in response to light and nitrogen levels. Plant Cell Tissue Organ Cult. 2018, 134, 29–39. [Google Scholar] [CrossRef]
  88. Yousaf, R.; Khan, M.A.; Ullah, N.; Khan, I.; Hayat, O.; Shehzad, M.A.; Khan, I.; Taj, F.; Din, N.U.; Khan, A.; et al. Biosynthesis of anti-leishmanial natural products in callus cultures of Artemisia Scoparia. Artif. Cells Nanomed. Biotechnol. 2019, 47, 1122–1131. [Google Scholar] [CrossRef] [Green Version]
Processes 10 01439 g001 550
Figure 1. Chemical structure of achillin compound.
Figure 1. Chemical structure of achillin compound.
Processes 10 01439 g001
Processes 10 01439 g002 550
Figure 2. Morphogenetic responses of leaf explants after 15 days of their incubation: (a) KIN (0.1 mg/L) with PIC (1.0 mg/L), (b) BAP (0.1 mg/L) with PIC (1.0 mg/L), (c) BAP (0.1 mg/L) with 2,4-D (0.1 mg/L), (d) KIN (0.1 mg/L) with NAA (1.0 mg/L), (e) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (f) BAP (0.1 mg/L) with NAA (1.0 mg/L). Scale bar = 10 mm.
Figure 2. Morphogenetic responses of leaf explants after 15 days of their incubation: (a) KIN (0.1 mg/L) with PIC (1.0 mg/L), (b) BAP (0.1 mg/L) with PIC (1.0 mg/L), (c) BAP (0.1 mg/L) with 2,4-D (0.1 mg/L), (d) KIN (0.1 mg/L) with NAA (1.0 mg/L), (e) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (f) BAP (0.1 mg/L) with NAA (1.0 mg/L). Scale bar = 10 mm.
Processes 10 01439 g002
Processes 10 01439 g003 550
Figure 3. Morphogenetic responses of nodal explants after 15 days of their incubation: (a) BAP (0.1 mg/L) with PIC (1.0 mg/L), (b) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (c) BAP (0.1 mg/L) with PIC (0.1 mg/L), (d) KIN (0.1 mg/L) with PIC (1.0 mg/L), (e) BAP (0.1 mg/L) with 2,4-D (0.1 mg/L), (f) BAP (0.1 mg/L with IAA (1.0 mg/L). Scale bar = 10 mm.
Figure 3. Morphogenetic responses of nodal explants after 15 days of their incubation: (a) BAP (0.1 mg/L) with PIC (1.0 mg/L), (b) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (c) BAP (0.1 mg/L) with PIC (0.1 mg/L), (d) KIN (0.1 mg/L) with PIC (1.0 mg/L), (e) BAP (0.1 mg/L) with 2,4-D (0.1 mg/L), (f) BAP (0.1 mg/L with IAA (1.0 mg/L). Scale bar = 10 mm.
Processes 10 01439 g003
Processes 10 01439 g004 550
Figure 4. Morphogenetic responses of internode explants after 15 days of their incubation: (a) BAP (0.1 mg/L) with NAA (1.0 mg/L), (b) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (c) BAP (0.1 mg/L) with IAA (0.1 mg/L), (d) BAP (0.1 mg/L) with PIC (0.1 mg/L). Scale bar = 10 mm.
Figure 4. Morphogenetic responses of internode explants after 15 days of their incubation: (a) BAP (0.1 mg/L) with NAA (1.0 mg/L), (b) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (c) BAP (0.1 mg/L) with IAA (0.1 mg/L), (d) BAP (0.1 mg/L) with PIC (0.1 mg/L). Scale bar = 10 mm.
Processes 10 01439 g004
Processes 10 01439 g005 550
Figure 5. Morphogenetic responses of root explants after 10 days of their incubation: (a) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (b) KIN (0.1 mg/L) with NAA (1.0 mg/L), (c) KIN (0.1 mg/L) with 2,4-D (0.1 mg/L). Scale bar = 10 mm.
Figure 5. Morphogenetic responses of root explants after 10 days of their incubation: (a) KIN (0.1 mg/L) with 2,4-D (1.0 mg/L), (b) KIN (0.1 mg/L) with NAA (1.0 mg/L), (c) KIN (0.1 mg/L) with 2,4-D (0.1 mg/L). Scale bar = 10 mm.
Processes 10 01439 g005
Processes 10 01439 g006 550
Figure 6. Achillin analysis by gas chromatography-mass spectrometry: (a) achillin isolated from A. ludoviciana wild plant; (b) achillin fragmentation pattern; (c) chromatogram of a callus extract from nodal explant with BAP (0.1 mg/L) plus NAA (1.0 mg/L), 1041.15 ± 16.47 µg/g dry biomass achillin compound.
Figure 6. Achillin analysis by gas chromatography-mass spectrometry: (a) achillin isolated from A. ludoviciana wild plant; (b) achillin fragmentation pattern; (c) chromatogram of a callus extract from nodal explant with BAP (0.1 mg/L) plus NAA (1.0 mg/L), 1041.15 ± 16.47 µg/g dry biomass achillin compound.
Processes 10 01439 g006
Table
Table 1. Effect of plant growth regulators on callus induction of A. ludoviciana through different types of explants in two incubation conditions.
Table 1. Effect of plant growth regulators on callus induction of A. ludoviciana through different types of explants in two incubation conditions.
PGRs(mg/L)PhotoperiodDarkness
LeafInternodeNodesRootLeafInternodeNodesRoot
Control00 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
KIN0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
BAP0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
NAA0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
1.00 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
2,4-D0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
1.00 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
IAA0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
1.00 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
PIC0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
1.00 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
KIN/NAA0.1/0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.00 ± 0.00 e46.67 ± 2.89 b80 ± 10.00 b80.00 ± 5.00 b0 ± 0.00 d40 ± 5.00 c26.67 ± 2.89 c70 ± 5.00 b
KIN/2,4-D0.1/0.10 ± 0.00 e71 ± 7.64 a0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.050 ± 5.00 c45 ± 5.00 b81.67 ± 2.89 b100 ± 0.00 a70.00 ± 5.00 b66.67 ± 2.89 b0 ± 0.00 d98.33 ± 2.86 a
KIN/PIC0.1/0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.090 ± 5.0 a0 ± 0.00 c0 ± 0.00 d80.00 ± 5.00 b100 ± 0.00 a55 ± 5.00 b71.67 ± 5.77 ab93.33 ± 2.89 a
KIN/IAA0.1/0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.00 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
BAP/NAA0.1/0.175 ± 5.0 b0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.095 ± 5.0 a0 ± 0.00 c100 ± 0.00 a0 ± 0.00 c95 ± 5.00 a0 ± 0.00 d83.33 ± 7.64 a0 ± 0.00 c
BAP/2,4-D0.1/0.153.33 ± 5.77 c0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.016.67 ± 2.89 d31.67 ± 2.89 b33.33 ± 5.77 c0 ± 0.00 c58.33 ± 2.89 c33.33 ± 5.77 c30 ± 5.00 c0 ± 0.00 c
BAP/PIC0.1/0.116.67 ± 2.89 d40 ± 10.00 b0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.085 ± 5.0 ab83.33 ± 7.64 a81.67 ± 2.89 b80 ± 5.00 b80 ± 5.00 b85.00 ± 5.00 a60 ± 10.00 b0 ± 0.00 c
BAP/IAA0.1/0.10 ± 0.00 e0 ± 0.00 c0 ± 0.00 d0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
0.1/1.00 ± 0.00 e0 ± 0.00 c75 ± 5.00 b0 ± 0.00 c0 ± 0.00 d0 ± 0.00 d0 ± 0.00 d0 ± 0.00 c
PGR = Plant growth regulators. Data represent mean ± standard deviation. Values with the same letters in columns are not statistically different according to Tukey’s multiple-range test (p ≤ 0.05).
Table
Table 2. Effect of plant growth regulators on callus–root induction of A. ludoviciana through different types of explants in two incubation conditions.
Table 2. Effect of plant growth regulators on callus–root induction of A. ludoviciana through different types of explants in two incubation conditions.
PGRs(mg/L)PhotoperiodDarkness
LeafLeafRoot
Control00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
KIN0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
BAP0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
NAA0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
2,4-D0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
IAA0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
PIC0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
KIN/NAA0.1/0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
0.1/1.073.33 ± 5.77 b0 ± 0.00 c0 ± 0.00 b
KIN/2,4-D0.1/0.198.33 ± 2.89 a95 ± 5.00 a100 ± 0.00 a
0.1/1.0100 ± 0.00 a0 ± 0.00 c0 ± 0.00 b
KIN/PIC0.1/0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
0.1/1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
KIN/IAA0.1/0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
0.1/1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
BAP/NAA0.1/0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
0.1/1.00 ± 0.00 c96.67 ± 2.89 a0 ± 0.00 b
BAP/2,4-D0.1/0.10 ± 0.00 c23.33 ± 2.89 b0 ± 0.00 b
0.1/1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
BAP/PIC0.1/0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
0.1/1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
BAP/IAA0.1/0.10 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
0.1/1.00 ± 0.00 c0 ± 0.00 c0 ± 0.00 b
PGR = Plant growth regulators. Data represent mean ± standard deviation. Values with the same letters in columns are not statistically different according to Tukey’s multiple-range test (p ≤ 0.05).
Table
Table 3. Effect of plant growth regulators on seedlings regeneration of A. ludoviciana through internodal and nodal segments in two incubation conditions.
Table 3. Effect of plant growth regulators on seedlings regeneration of A. ludoviciana through internodal and nodal segments in two incubation conditions.
PGR(mg/L)NodesInternodes
PhotoperiodDarknessPhotoperiodDarkness
Control0.023.33 ± 5.77 c0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
KIN0.10 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
BAP0.10 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
NAA0.10 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
1.00 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
2,4-D0.10 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
1.00 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
PIC0.10 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
1.00 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
IAA0.10 ± 0.00 d0 ± 0.00 d 0 ± 0.00 e0 ± 0.00 b
1.00 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
KIN/NAA0.1/0.10 ± 0.00 d23.33 ± 2.87 c0 ± 0.00 e0 ± 0.00 b
0.1/1.00 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
KIN/2,4-D0.1/0.10 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
0.1/1.00 ± 0.00 d0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
KIN/PIC0.1/0.10 ± 0.00 d0 ± 0.00 d91.67 ± 2.89 a0 ± 0.00 b
0.1/1.095 ± 5.00 a0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
KIN/IAA0.1/0.10 0 ± 0.00 d23.33 ± 2.89 c0 ± 0.00 e0 ± 0.00 b
0.1/1.026.67 ± 2.89 c93.33 ± 2.89 a0 ± 0.00 e0 ± 0.00 b
BAP/NAA0.1/0.128.33 ± 2.89 c23.33 ± 2.89 c0 ± 0.00 e0 ± 0.00 b
0.1/1.041.67 ± 2.89 b0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
BAP/2,4-D0.1/0.196.67 ± 2.89 a33.33 ± 2.89 b0 ± 0.00 e0 ± 0.00 b
0.1/1.033.33 ± 2.89 bc0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
BAP/PIC0.1/0.10 ± 0.00 d0 ± 0.00 d16.67 ± 5.78 d0 ± 0.00 b
0.1/1.098.33 ± 2.89 a0 ± 0.00 d0 ± 0.00 e0 ± 0.00 b
BAP/IAA0.1/0.10 ± 0.00 d0 ± 0.00 d36.67 ± 5.78 c100 ± 0.00 a
0.1/1.091.67 ± 2.89 a0 ± 0.00 d53.33 ± 2.89 b0 ± 0.00 b
PGR = Plant growth regulators. Data represent mean ± standard deviation. Values with the same letters in columns are not statistically different according to Tukey’s multiple-range test (p ≤ 0.05).
Table
Table 4. Achillin content in the biomass of different morphogenetic responses (callus and plantlets) of A. ludoviciana analyzed by GC-MS.
Table 4. Achillin content in the biomass of different morphogenetic responses (callus and plantlets) of A. ludoviciana analyzed by GC-MS.
Plant ResourceMorphogenetic ResponseGrowing ConditionsPGRs (mg/L)Achillin
(µg/g Dry Biomass)
AuxinCytokinin
LeavesCallusPhotoperiodPIC (1.0)KIN (0.1)1703.05 ± 40.92
NodesCallusTotal darknessNAA (1.0)BAP (0.1)1041.15 ± 16.47
LeavesCallusPhotoperiod2,4-D (1.0)BAP (0.1)88.34 ± 8.15
RootsCallusTotal darkness2,4-D (1.0)BAP (0.1)12.79 ± 13.98
NodesPlantletsPhotoperiodPIC (0.1)BAP (0.1)1880.01 ± 42.67
InternodesPlantletsPhotoperiodPIC (0.1)KIN (0.1)334.03 ± 13.29
InternodesPlantletsPhotoperiodNAA (0.1)BAP (0.1)28.11 ± 1.03
InternodesPlantletsPhotoperiodPIC (0.1)BAP (0.1)1482.17 ± 53.44
InternodesPlantletsTotal darknessIAA (0.1)BAP (0.1)53.46 ± 12.95
InternodesPlantletsPhotoperiodIAA (0.1)BAP (0.1)1589.25 ± 30.42
PGR = plant growth regulators.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Sánchez-Ramos, M.; Berman-Bahena, S.; Alvarez, L.; Sánchez-Carranza, J.N.; Bernabé-Antonio, A.; Román-Guerrero, A.; Marquina-Bahena, S.; Cruz-Sosa, F. Effect of Plant Growth Regulators on Different Explants of Artemisia ludoviciana under Photoperiod and Darkness Conditions and Their Influence on Achillin Production. Processes 2022, 10, 1439. https://doi.org/10.3390/pr10081439

AMA Style

Sánchez-Ramos M, Berman-Bahena S, Alvarez L, Sánchez-Carranza JN, Bernabé-Antonio A, Román-Guerrero A, Marquina-Bahena S, Cruz-Sosa F. Effect of Plant Growth Regulators on Different Explants of Artemisia ludoviciana under Photoperiod and Darkness Conditions and Their Influence on Achillin Production. Processes. 2022; 10(8):1439. https://doi.org/10.3390/pr10081439

Chicago/Turabian Style

Sánchez-Ramos, Mariana, Samantha Berman-Bahena, Laura Alvarez, Jessica Nayelli Sánchez-Carranza, Antonio Bernabé-Antonio, Angélica Román-Guerrero, Silvia Marquina-Bahena, and Francisco Cruz-Sosa. 2022. "Effect of Plant Growth Regulators on Different Explants of Artemisia ludoviciana under Photoperiod and Darkness Conditions and Their Influence on Achillin Production" Processes 10, no. 8: 1439. https://doi.org/10.3390/pr10081439

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop