1. Introduction
The savin juniper, or savin (Juniperus sabina L. var. sabina; Cupressaceae), is a juniper species native to the mountains of Central and Southern Europe and Western and Central Asia. It typically thrives at altitudes ranging from 700 to 3300 m [1]. The genus Juniperus is of particular importance in arid and semi-arid areas because it is well adapted to severe drought and poor soil conditions [2]. The resilience and hardiness of junipers make them suitable for reforestation in less productive or degraded areas [1]. The Juniperus genus also includes important groups of species from which to obtain EOs because the number of aromatic elements is very high [3,4,5,6,7].
In recent times, there has been an increasing interest in the extraction of essential oils (EOs) from forest species, propelled by their substantial applications across diverse fields [8,9,10,11]. For this reason, essential oil extraction procedures have been recently included in forest management planning—for example, to obtain compounds with antibacterial properties in coniferous forests [12]. Many fragrances incorporate the oils of aromatic forest plants—for instance, lavender oil and rosemary oil [10]. Moreover, plant EOs used singly or in combination can be effective tools to inactivate microorganisms and microbial agents [4,12].
The increased interest in obtaining EOs from forest species represents an opportunity to utilize the great potential of junipers regarding the foliar metabolites. In particular, Juniperus sabina (J. sabina) is one of the most valuable conifers for essential oil production for use in industry and medicine. Sabinene has often been characterized as the predominant constituent in the essential oil extracted from the leaves of J. sabina [13,14,15,16,17,18], with values of this compound between 24 and 61%. Other EOs included in the leaves are α-pinene, cedrol, limonene, and terpinen-4-ol [13]. In other juniper species, sabinene is also an important component—for example, in Juniperus communis [19] or Juniperus indica [20].
Leveraging its distinct and abundant phytochemical composition, the EOs derived from J. sabina have found applications in both cosmetics and pharmacology [3,15,16,21]. Additionally, studies have demonstrated that the antibacterial and antimicrobial activities of the essential oils present in the leaves of J. sabina [5], as well as antioxidant properties [22]. The essential oil of J. sabina is toxic for some insects and can be used as a natural pesticide [23]. Because the leaves of savin also contain podophyllotoxin, this species has recently been used to obtain compounds as precursors for anticancer drugs [5].
The production of EOs is subject to diverse factors, including the growing environment, altitude, latitude, harvesting time, and the specific plant organ employed for oil extraction [24,25]. Farhi et al. [15] also noted that the essential oil yields were higher for male (0.86%) sabin individuals compared to female (0.65%). Additional problems regarding the use of juniper leaves in forest management are the random distribution of individuals within coniferous forests and the difficulties in their natural regeneration. Seed dormancy, the low viability of embryos, and poor site quality are factors that condition recruitment [1]. These limitations in forest management mean that junipers are frequently protected by nature conservation legislation, which also limits their use. Although the seeds of this species have been used for propagation, it is necessary to use different treatments that require long periods of time to alleviate seed dormancy [1].
Under these circumstances, vegetative propagation by cuttings is recommended for junipers [1] because the production of cuttings in nurseries permits the regular production of both plants and EOs, independently of site and legislation factors. In the nursery, the essential oil evaluation can also be performed in a second phase using the leaves of cuttings based on the hydrodistillation method [8,26]. Finally, the type and percentage of individual components of the EOs of samples can be measured by gas chromatography–mass spectrometry analysis (GC-MS analysis) [27,28,29]. As occurs for the essential oil composition, the harvesting time plays also a very important role in the root development success [30], as well as selecting an adequate substrate for rooting [31].
To optimize cutting production, it is commonly observed that treating stem cuttings with varying hormone concentrations in a suitable substrate (rooting medium) is necessary to enhance rooting [31,32,33]. Propagating junipers through cuttings has shown successful improvements in rooting percentage by pretreating with indole butyric acid in a perlite-based substrate [34]. However, natural compounds can be used instead of chemical growth regulators (such as IBA treatment) to maximize the rooting performance because they are sustainable compounds. Coconut juice stands as an alternative and natural hormone that may facilitate an increase in cutting rooting, owing to the presence of essential hormones such as auxin, cytokinin, and gibberellin [35,36].
Despite the limited research conducted to date, coconut juice has demonstrated success in promoting the rooting of forest species [36,37,38]. For example, in Parkia biglobosa, it resulted in an increased percentage of rooted cuttings and the formation of calluses [36]. Additionally, in Cordia millenii, Enantia chlorantha, and Celtis durandii, coconut juice facilitated rooting percentages of 60%, 50%, and 40%, respectively [39]. In a study of Agele et al. [37], the treatment with coconut juice demonstrated superiority over IBA and IAA in terms of bud retention, rooting, leaf development, and the overall survival of plantlets in four African species. Nevertheless, there is no existing demonstration of the application of coconut juice for plant rooting in coniferous species. Furthermore, there is a notable absence of research where coconut juice pretreatment has been employed for juniper cuttings. Growth regulators have the potential to simultaneously increase essential oil yield by influencing the metabolic pathways associated with essential oil biosynthesis [38].
As a result, our experimental layout was designed to explore an efficient pretreatment method (using coconut juice in a suitable substrate) for the vegetative propagation of savin juniper through stem cuttings. Additionally, our objective was to enhance the essential oil production from the leaves. Thus, the main aims of this study were (i) to study the rooting performance in J. sabina cuttings subjected to four levels of coconut juice as pretreatment (25%, 50%, 75%, and 100%), as well as using four substrates (perlite, mixed substrate, pumice, and perlite–cocopeat), (ii) to examine the production of essential oil in the leaves of the cuttings as an outcome of the applied treatments, and (iii) to evaluate the effects of the harvesting time (growing season) on rooting and essential oil production in leaves. The results shed light on the potential development of forest biotechnology to produce cuttings and EOs from juniper species.
2. Materials and Methods
2.1. Sampling of Cuttings, Pretreatment of Cuttings with Coconut Juice, and Substrate Preparation
The J. sabina cuttings were collected from their natural habitat in the Chahar-bagh mountains of Gorgan, located in northern Iran (Figure 1). This region represents one of the primary Mediterranean populations at a higher altitude, approximately 2700 m above sea level. Based on the 30-year average values, the mean annual temperature at the site was 9.2 °C, and the mean annual precipitation was 429 mm. Temperature variations between summer and winter ranged from 23 °C to −5 °C (data obtained from the Gorgan climatic station at 46°06 N, 28°00 W; 2600 m.a.s.l.). The crowns were approximately 2 × 2 m in length and width. The ring diameter of shrubs was 20.0 cm on average, and the height was 1.5 m (referring to old and horizontal shrubs). Because male individuals generally have greater yields of EOs [15], a total of 20 male shrubs were used for this experiment, and they were all grown in the same area, with the same ecological environment. The experiment was conducted in the winter, spring, summer, and fall of 2017. Stem cuttings were exclusively collected from the upper crowns of male trees.
The cuttings were harvested in the morning. Subsequently, the stem cuttings were prepared to be 15 cm in length and 0.5–0.7 cm in diameter [41] for treatment and cultivation in a greenhouse. We used hardwood or semi-hardwood (in summer) cuttings because the juniper plant material was taken from the tips of the branches, and it included the previous year’s growth [31]. The cuttings were placed in a greenhouse with an automatic system of micro-irrigation and bottom heat. The mean daily temperature during the experiment was 22 °C, and the mean relative humidity was 77%. The light entering the greenhouse varied based on the growing season.
For the treatment of stem cuttings, four doses of coconut juice were used. Thus, the levels analyzed were 0% (control), 25%, 50%, 75%, and 100% of coconut juice. To prepare each dose, the percentage of coconut juice was placed in the container and then the volume was increased to 100% using distilled water. Distilled water was not used in the case of 100% coconut juice. The number of cuttings required for each treatment was separated and placed in a container containing the desired amount of coconut juice for 24 h in a greenhouse. In the 0% treatment, only distilled water was used. Following this period, the cuttings were planted in the four designated substrates (each approximately 10 kg): (i) perlite; (ii) mixed rooting substrate (composed of sand (20%), perlite (20%), cocopeat (20%), vermicompost (20%), and potash (20%)); (iii) perlite–cocopeat (1:1); and (iv) mineral pumice. For each combination of coconut juice pretreatment and substrate, three replicates were prepared, with nine cuttings per biological replicate. Thus, a total of 540 cuttings in each growing season were cultivated.
2.2. Rooting Percentages and Chemical Compounds of Cuttings: Essential Oil Evaluation
To assess the rooting percentage of each treatment, the roots were extracted from all rooted cuttings in each treatment. The experiment was replicated three times, with each replication comprising 9 cuttings, resulting in a total of 27 cuttings.
For essential oil extraction, leaf samples were subjected to the hydrodistillation method [26]. Based on the literature, hydrodistillation stands out as the most frequently employed method for essential oil extraction including leaves [42]. This method has been successfully applied to juniper plant material [43]. The widespread adoption of the hydrodistillation method is attributed to its simplicity in operation and maintenance, as well as its cost-effectiveness when compared to alternative methods [44] such as solvent extraction, fluid extraction, and steam distillation. Additionally, hydrodistillation utilizes water as the extraction medium, operating at lower temperatures than other methods; this proves advantageous for preserving heat-sensitive compounds in EOs, maintaining the natural characteristics of the oil [45]. As a result, the method is scalable for large-scale production, rendering it applicable for industries engaged in the commercial production of EOs.
To initiate the hydrodistillation procedure, leaf samples were dried in the shade until they attained a constant weight [46]. Subsequently, 50 g of leaves per treatment were weighed and placed in 1 L balloon flasks, followed by the addition of 700 mL of distillation water. The sample underwent water distillation in a Clevenger apparatus at 250 °C for 3 h. This process was conducted under atmospheric pressure using a hydrodistillation apparatus equipped with an electrical resistance heater for water heating and boiling. The prepared leaf suspension was introduced into the distillation flask, and heating commenced. Following this stage, the EOs emerged in the upper part of the balloon and were separated from the water through decantation. The EOs obtained through distillation were dehydrated and subsequently stored at 4 °C in dark glass bottles until subjected to analysis. To calculate the percentage of essential oil, the weight of the obtained essential oil was divided by the weight of the plant sample utilized for extraction [12]: EO (%) = weight of obtained EO (mL)/weight of plant sample (kg).
The oil underwent analysis through an analytical GC-MS method. Gas chromatography combined with mass spectrometry is widely regarded as the most effective method for separating and detecting EOs [29]. GC-MS analysis permits the identification of phytochemical compounds based on the peak area, retention time, molecular weight, and molecular formula [27,28]. For the optimization process, a gas chromatograph (Shimadzu GC-2010 Plus AF, Tokyo, Japan) equipped with a split/splitless injection port and a flame ionization detection system was utilized. The initial column temperature was 40 °C. It was increased to 100 °C at a rate of 20 °C min−1, and then raised to 220 °C at 3 °C min−1. Subsequently, the temperature was increased to 280 °C at a rate of 30 °C min−1, with a hold time of 2 min. To separate the desorbed polycyclic aromatic hydrocarbons (PAHs) from the needle trap desorption (NTD), the injection was performed using split mode at an injection port temperature of 280 °C. The detector temperature was set at 280 °C.
It is important to note that at the onset of each season, the type and percentage of individual components in the essential oil samples were measured using the GC-MS procedure. Samples were fresh cuttings from the first stage of each season, from which their essential oil content was measured, without any auxin or coconut extract treatment. At the first stage of each season, there was one extraction of EOs, as shown in Figure 2.
2.3. Data Analysis
A factorial arrangement of treatments [47] was employed to analyze the effects of three main factors (dose of coconut juice, season, and substrate) on two dependent variables (rooting performance, as % of rooting, and essential oil in leaves, in %). The analyzed factors were (i) the concentration of coconut juice (five levels: 0% or control, 25%, 50%, 75%, and 100%), (ii) the growing season (four levels: winter, spring, summer, and fall, corresponding to the sampling season and with a difference of 3 months between treatments), and (iii) the substrate (four levels: perlite, perlite–cocopeat, pumice, and mixed rooting substrate). This represented a 5 × 4 × 4 factorial with 80 combinations of factor levels or treatments. Three replicates (9 cuttings each) were considered for each treatment for statistical analysis. Thus, the sample in each season was 5 doses of coconut juice × 4 substrates × 3 replicates × 9 cuttings/replicate = 540 cuttings.
An additional treatment level was introduced for the dependent variable related to EOs: the percentage of essential oil in fresh samples (stem cuttings that were not planted and were prepared at the beginning of each season). This addition aimed to facilitate the comparison of treatment effects between nontreated cuttings (at the beginning of each season) and treated cuttings (at the end of each season).
The SAS® statistical software (SAS Institute Inc., Cary, NC, USA) was used to detect significant factors and to evaluate significant differences between mean values in the levels of treatment. The means were compared using the PROC GLM procedure in SAS, employing multifactor analysis of variance (a three-way ANOVA model) with a significance level of 5% (p < 0.05). Seasonal analysis was conducted through a two-way ANOVA, with the exclusion of season as a primary factor in the overall model. Independent ANOVAs were chosen for this study, avoiding a mixed-design or repeated-measures ANOVA, as the measurements were independent (distinct stem cuttings were used for each treatment and season).
A Fisher’s least significant difference (LSD) test (p < 0.05) was used to explore the significant differences between treatments [48]. To employ this statistical method effectively, it is preferable for the data to exhibit a normal distribution. However, this is not applicable to proportions, as their values typically range between zero and one. Furthermore, errors should demonstrate independence and follow a normal distribution with consistent variance. To meet these assumptions, a logarithmic transformation was applied. Specifically, for variables such as the percentage of rooting and essential oil content, the analysis involved [ln (r + 0.5)], where ‘r’ represented the percentage (divided by 100). Given that this transformation necessitates numerical data above zero, a small value (0.5) was added to the variable before conducting the transformation.
3. Results
3.1. Multifactor ANOVA Results
Table 1 indicates that the rooting percentage and essential oil percentage were significantly influenced by the cutting pretreatment, season, and substrate (p < 0.05). Thus, the results confirmed the strong seasonality (effect of harvesting time) of the development and essential oil production of stem cuttings pretreated with coconut juice. Regarding second-order interactions, the coconut juice treatment × season interaction significantly affected both the % rooting and the essential oil percentage, thus confirming the seasonality (Table 1). We highlight that the coconut juice treatment × substrate interaction was also significant for the dependent variables (rooting and EOs) (p < 0.05; Table 1). The effect of the sampling time on the dependent variables was also manifested as several significant effects within seasons, but only for the pretreatment with coconut juice in cuttings.
3.2. Rooting Performance Using Coconut Juice
As confirmed in the multifactorial ANOVA, the pretreatment with coconut juice significantly increased the rooting performance compared to the control treatment in all growing seasons (Figure 3a). Moreover, the percentage dose of coconut juice within seasons showed several significant effects. The highest root percentage was seen in spring at 25% coconut juice with 30.66 ± 0.52% rooting (mean ± standard error; within and between seasons; p < 0.05). Using the treatment with 50% coconut juice, rooting of 23.35 ± 0.52% was noted in spring. Slight rooting was noted in the treatment involving 75% coconut juice in autumn (11.11 ± 1.04). In contrast, in winter, no rooting was observed at any of the coconut juice levels (Figure 3a). In consequence, our results showed that it is preferable to root juniper cuttings in spring, adding a dose of 25% coconut juice.
Analyzing the interaction between coconut juice and the substrates (Figure 3b), the highest rooting was seen in the substrates of perlite–cocopeat (at 25% coconut juice; 37.10 ± 1.04%) and pumice (at 50% coconut juice; 37.01 ± 1.04%). For the perlite and mixed substrates, a higher % of rooting was also noted for the level of 25% coconut juice (29.21% and 33.11%, respectively). In general, the control showed a % of rooting below 20%, except for the mixed substrate. Therefore, pretreatment with 25%–50% coconut juice amplified the impact of the substrate on the rooting of cuttings.
3.3. Production and Percentage of EOs with Coconut Juice as Pretreatment
Analyzing the effects of the levels of coconut juice by season on the percentage of essential oil obtained from cuttings (Figure 4a), we verified that this pretreatment significantly improved the percentage of EOs at 50% in spring (0.82 ± 0.16; mean ± standard error). This value was not statistically significant between seasons because similar percentages of essential oil were obtained for fresh samples in winter and fall (1.08 ± 0.11% and 0.85 ± 0.15%, respectively; mean ± standard error).
In this sense, the highest quantity of essential oil was observed in the fresh sample during the winter (Figure 4a), probably because, during the maintenance of cuttings in winter, the amount of essential oil was significantly reduced. This tendency was also noted in the fall. In this season, the largest amount of essential oil in pretreated cuttings was seen under the treatment with 75% coconut juice (0.52 ± 0.16%; mean ± standard error).
The influence of substrates on the percentage of EOs with varying concentrations of coconut juice in the cuttings (Figure 4b) indicated that the mixed rooting substrate, pumice and perlite–cocopeat, produced the largest amount of essential oil in the fresh samples, and, during the maintenance of the cuttings in the substrate, the percentage of essential oil decreased. Nevertheless, the highest percentage of EOs among substrates was noted in perlite (which was the lightest substrate) with a 25% concentration of coconut juice (1.16 ± 0.19%, significantly higher than the rest of the treatments, p < 0.05), whereas lower content of essential oil was observed in the mixed rooting substrate (which was the heaviest substrate).
3.4. Chromatogram GC-MS Analysis and Individual Components of Essential Oil
The chromatogram obtained through GC-MS analysis of the EOs extracted from J. sabina revealed the presence of numerous peaks in each season (Figure 5). To ascertain the seasonality in essential oil production, the components corresponding to these peaks were identified and documented (Table 2).
Table 2 presents the assessment of the individual components of the essential oils (EOs) obtained from the samples at the commencement of each season (the table includes individual components extracted in each season). The values are based on the percentage and type of the various obtained compounds.
As can be seen from Table 2, the individual components of the EOs of J. sabina cuttings also changed as a function of the season. In winter, the largest number of total components was obtained from sabinene (31.22%), but in spring, summer, and fall, it was obtained for nerolidol (8.5%), 3-carene (16.8%), and beta-pinene (34.7%), respectively. The component terpinen-4-ol was obtained in all seasons except in spring. Therefore, the type and percentage of each oil compound showed high seasonality.
4. Discussion
4.1. Rooting of the Cuttings with Coconut Juice as Pretreatment
In our study, the highest rooting percentage was obtained with a level of 25% coconut juice. This natural pretreatment improved the rooting performance compared with nontreated plant material in all growing seasons. The positive effect of coconut juice on the rooting percentage in cuttings of savin juniper could be due to the inclusion of growth regulators such as auxin and cytokinin in its composition [33,36,49]. The components of coconut juice also include sugar, amino acids, mayo-inocythol, and phenyl-urea, which increase cytokinin activity, and it also contains indole acetic acid and gibberellic acid [50]. In general, plant growth regulators stimulate mainly auxiliary root formation because their components induce adventitious roots [36,51]. For these reasons, the use of appropriate coconut juice levels for cuttings as a function of each species can increase the level of rooting.
The use of IBA has also shown a greater effect on rooting than coconut juice in other plants [52,53]. However, IBA is a chemical compound that, under certain doses, can damage cuttings [6]. Therefore, the use of coconut juice has been recommended to increase the rooting of cuttings, rather than other treatments [49].
The timing of the cutting also plays a crucial role in the success of rooting. In this context, our findings indicate that the optimal time for preparing cuttings with coconut juice was during the spring season. In this season, a higher percentage of nerolidol (8.58%) was also obtained. Our results align with the findings of Fragoso et al. [54] and Tektas et al. [55], both of whom identified spring as the optimal season for the rooting of Juniperus cuttings. However, other authors have published different results. For example Chowdhuri [56] demonstrated that summer was the optimal season for rooting in Juniperus chinensis, while Ali ahmad korouri et al. [25] studied Juniperus horizontalis, whose cuttings were most rooted when they were prepared between November and February. Although many species are best rooted before the wood has hardened (early winter or late spring), other species show the best rooting in the following seasons, when the plant physiology is more active [31,57].
Referring to the substrate, in our study, the percentage of rooting was improved in the substrates of perlite–cocopeat (at 25% coconut juice; 37.10%) and pumice (at 50% coconut juice; 37.01%). On the contrary, in a study of Juniperus procumbens, the most favorable substrates were vermiculite and perlite [33]. The results of our study also differ from the findings obtained by Khoushnevis et al. [58], with 28% of rooting in Juniperus oblonga using fine and harsh bed, and Stuepp et al. [59], with 16% of rooting in Juniperus chinensis using vermiculite. The perlite–cocopeat substrate maintains suitable moisture to prevent the cutting ends from drying out and to provide sufficient air to facilitate rooting and prevent disease spread at the base of the cuttings [31,60]. It is likely that there is an optimum temperature for the substrate to facilitate root formation, as growth and rooting cannot occur at low temperatures (or occurs very slowly).
4.2. Effects of Pretreatment with Coconut Juice on Essential Oil Production
Our findings showed that in spring, the pretreatment with 50% coconut juice significantly increased the EOs (0.82%) when compared with the control or fresh samples. The obtained yield was within the interval reported in the literature for the production of EOs from J. sabina leaves, which varied between 0.6 and 1.9% [13,15,61]. The essential oil content can be influenced by both the timing of exogenous application and the specific type of growth regulator used [62]. In a broad sense, plant growth regulators influence the production of EOs by impacting various aspects of plant growth, including the biomass of leaves, fruits, or flowers, as well as essential oil biosynthesis [38]. In particular, coconut juice has the additional effects of increasing the roots and enabling the greater production of calluses in the cuttings, which is due to the presence of auxins and cytokinin in this juice [36].
The dominance of root growth over shoot growth has been associated with variations in sesquiterpenoid quantities and the preferential direction of biosynthesis toward monoterpenoids [63]. This relationship could explain in part the low values of the sesquiterpene nerolidol (8.58%) extracted as the principal component in spring, when the effect of the coconut juice was at its maximum with regard to both rooting performance and essential oil production. In contrast, when the coconut juice did not affect the EOs and the rooting performance was very low or zero (in fall and winter, respectively), the dominant components of the EOs in J. sabina leaves were generally the monoterpenes sabinene (31.22%, in winter) and beta-pinene (34.76%, in fall). Growth regulators can also play a role in shaping storage structures, a decisive factor in terpene biosynthesis [38]. For example, the positive impact of plant hormone application on structure formation was observed in a study of cytokinins in Thymus mastichina, influencing essential oil production [64].
On the other hand, when cuttings are cultivated ex vitro, as in the current study, additional factors exert a significant influence on the chemical composition of the EOs. It is recognized that the main components of the essential oil are principally determined genetically, but the compounds may vary under different edaphic and climatic conditions [15,65]. Moreover, growth regulators not only influence plant growth, but affect biochemical processes and gene regulation as well [66]. As a consequence, both the timing of exogenous application and the type of growth regulator can influence the composition of EOs [62].
4.3. Components of Essential Oil in J. sabina as a Function of Growing Season
As occurred for the total yield of EOs, our findings revealed a notable seasonal pattern in the production of various components of the EOs in J. sabina, which is consistent with the earlier findings reported by Adams et al. [6], Ali Ahmad Korouri et al. [25], and Salehi Shenjani and Mirza [67].
As noted above, the metabolic processes of EOs reflect the interactions between plants and the environment [68,69], and factors such as soil composition and precipitation levels play a crucial role in shaping distinct structures in plants [70], thus developing distinct chemotypes [1]. In this context, the essential oil yield and composition can also be affected by the distribution of precipitation throughout the year [43]. This observation could elucidate the pronounced seasonality in the production of both the overall essential oil and the various individual components extracted.
As a result of this seasonality, the most important components extracted by season were beta-pinene in fall (34.7%) and sabinene in winter (31.2%). In general, previous studies have reported similar percentages of sabinene from J. sabina leaves [14,16,31]. Sabinene, a significant naturally occurring bicyclic monoterpene, finds applications in flavorings, perfume additives, fine chemicals, and advanced biofuels [8]. Different values of sabinene were reported by Semerdjieva et al. [18], with 61% (the highest percentage included in the literature), and Farhi et al. [15], with 16.69% in female plants. The large differences found in the percentages of sabinene in J. sabina confirm the relationship between EOs and site factors. As another, similar example in the Juniperus genus, the leaves of Juniperus communis that produced sabinene were found only in three localities from 34 habitats, beside plants with needles that biosynthesized α-pinene-chemotype oils [71].
In contrast, beta-pinene, the primary constituent of the oil extracted from leaves in autumn, was found to be absent in other studies. In earlier studies, it was established that α-pinene is a noteworthy constituent of the EOs in J. sabina [23,67], as in the study of Adams et al. [8], where the main components of Juniperus excelsa and Juniperus polycarpos were alpha-pinene and limonene. In contrast, in the study of Ghorbanzadeh et al. [72], high levels of myrtenyl acetate were detected in J. sabina tissue; however, this terpenoid was absent in both of the other studied species and has not been reported in other studies focusing on juniper. The study of Zhang et al. [7] in Juniperus chinensis showed that the major constituent was beta-phellandrene (26.64%–39.26%), followed by terpinen-4-ol (6.53%–11.89%). Therefore, the type of compound and its importance in the total oil produced varies greatly depending on the site.
Beta-pinene, as emphasized in the literature, exhibits parameters relevant to neurological treatment. It is recommended that future research explores the therapeutic potential of terpenes for conditions such as stroke, ischemia, inflammatory and neuropathic pain (including migraine), cognitive impairment (associated with Alzheimer’s disease and aging), insomnia, anxiety, and depression [73].
Finally, 3-carene (the principal compound in summer) is an antimicrobial monoterpene and it has antibacterial activity [74], whereas nerolidol, which is a major component in spring, has received approval from the US Food and Drug Administration as a food-flavoring agent and has exhibited antimalarial activity [75].
Therefore, the type and percentage of each compound varies at different harvesting times, and these results can be applied to select the appropriate rooting season based on the individual components that are necessary.
5. Conclusions
Our study has demonstrated the effectiveness of using coconut juice to enhance the rooting performance of J. sabina cuttings, mainly in spring. In spring, the treatment with coconut juice also significantly enhanced the essential oil of juniper leaves.
Among the essential oil components previously cited, sabinene has been confirmed as the major compound, specifically during the winter season. However, in spring, summer, and fall, the main compounds in the leaves of J. sabina were nerolidol, 3-carene, and beta-pinene, respectively. In particular, pretreatment with coconut juice in spring would maximize the production of nerolidol, which has been recently cited as a flavoring and antimalarial compound.
Finally, further studies on the effects of coconut juice on both rooting and essential oil content in other juniper or conifer species should be carried out, since it is a natural, sustainable compound, and its use permits an enhancement in the rooting and production of EOs using natural sources. Unresolved issues that remain to be studied include the utilization of different doses of coconut juice and different types of cuttings (comparing green and woody cuttings) and substrates to improve the efficiency of this method in coniferous species, thus overcoming the limitations of chemical treatments. In addition, the period of study can be extended to elucidate the changes that occur as the plants age.
Conceptualization, M.A., B.Z., H.Z. and A.R.N.; formal analysis, M.A., F.A.G.-M. and A.R.N.; investigation, M.A. and F.A.G.-M.; methodology, M.A. and H.Z.; project administration, H.Z.; resources, B.Z.; supervision, F.A.G.-M., B.Z., H.Z. and A.R.N.; validation, A.R.N.; writing—original draft, M.A.; writing—review and editing, F.A.G.-M. and H.Z. All authors have read and agreed to the published version of the manuscript.
Not applicable.
The data presented in this study are available on request from the corresponding author. The data are not publicly available due to privacy.
This study was part of a PhD thesis supported by the Lorestan University (Iran) and a grant for a scientific stay by Maliheh Abshahi at the University of Castilla-La Mancha (Spain), supported by the Ministry of Science, Research and Technology, Iran. We are grateful to four anonymous reviewers for their comments, which allowed us to improve the initial manuscript.
The authors declare no conflict of interest.
Footnotes
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![View Image - Figure 1. The global distribution of various populations of J. sabina L. depicted in grey. Reproduced from Adams et al. [40]. Open access by the College of Health Professions at ScholarWorks @ University of Texas Rio Grande Valley (UTRGV). The specific sampling area for stem cuttings is highlighted with a red circle.](https://media.proquest.com/media/hms/THUM/2/x4CBX?_a=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%2BgIBWYIDA1dlYooDHENJRDoyMDI1MDcwMzA4MTExMzg3MDo0MTA2MjA%3D&_s=2Ip3Zq9ypDX15teab45GKjVWScI%3D "View Image - Figure 1. The global distribution of various populations of J. sabina L. depicted in grey. Reproduced from Adams et al. [40]. Open access by the College of Health Professions at ScholarWorks @ University of Texas Rio Grande Valley (UTRGV). The specific sampling area for stem cuttings is highlighted with a red circle.")
Enlarge this image.Figure 1. The global distribution of various populations of J. sabina L. depicted in grey. Reproduced from Adams et al. [40]. Open access by the College of Health Professions at ScholarWorks @ University of Texas Rio Grande Valley (UTRGV). The specific sampling area for stem cuttings is highlighted with a red circle.
Enlarge this image.Figure 2. Measurement of plant EOs in cuttings of J. sabina: (a) experimental plot with savin juniper sampled, (b) unrooted vs. rooted cuttings that were treated with coconut juice as a plant growth regulator (the effectiveness of the rooting pretreatment can be observed), (c) the bench of the greenhouse with cuttings cultured in substrates, and (d) Clevenger apparatus used to measure percentage of essential oil.
Enlarge this image.Figure 3. Average percentages of rooting for the treatments involving coconut juice (a) within seasons and (b) as a function of substrate. Distinct lowercase letters denote statistically significant differences (p ≤ 0.05; LSD test) between the treatments for each (a) season or (b) substrate. Different uppercase letters signify statistically significant differences (p ≤ 0.05; LSD test) between (a) seasons or (b) substrates for the treatments (only treatments with rooting above 20% are differentiated in significant groups; horizontal line). Sample data = 540 cuttings for each season and substrate. For treatments not represented in the figure, no cuttings were rooted. Error bars: standard error. C: control (nonpretreated cuttings).
Enlarge this image.Figure 4. Mean internal essential oil values for the different treatments with coconut juice (a) within seasons and (b) within substrates. Distinct lowercase letters denote statistically significant differences (p ≤ 0.05; LSD test) between the treatments for each (a) season or (b) substrate. Different uppercase letters signify statistically significant differences (p ≤ 0.05; LSD test) between (a) seasons or (b) substrates for the treatments (only treatments with EOs content above 0.60% are differentiated in significant groups; horizontal line). A total of 540 stem cuttings in each season and substrate were treated. Treatments in summer are not represented in the figure (due to the destruction of most cuttings during their maintenance in the substrate, no samples remained to measure the essential oil). Error bars: LSD intervals. FS: fresh sample; C: control (nonpretreated cuttings).
Enlarge this image.Figure 4. Mean internal essential oil values for the different treatments with coconut juice (a) within seasons and (b) within substrates. Distinct lowercase letters denote statistically significant differences (p ≤ 0.05; LSD test) between the treatments for each (a) season or (b) substrate. Different uppercase letters signify statistically significant differences (p ≤ 0.05; LSD test) between (a) seasons or (b) substrates for the treatments (only treatments with EOs content above 0.60% are differentiated in significant groups; horizontal line). A total of 540 stem cuttings in each season and substrate were treated. Treatments in summer are not represented in the figure (due to the destruction of most cuttings during their maintenance in the substrate, no samples remained to measure the essential oil). Error bars: LSD intervals. FS: fresh sample; C: control (nonpretreated cuttings).
Enlarge this image.Figure 5. GC chromatograms of essential oil of J. sabina in each season. The presence of several peaks in each season permitted the components of the essential oil to be determined.
Results of the multifactor ANOVA examining the influence of the main factors on rooting performance and essential oil content of cuttings across the four seasons. The table displays p-values for the three primary effects (pretreatment, substrate, and season) and their two-way interactions. Additionally, it includes p-values for effects within each season (pretreatment, substrate, and their interaction). Treatments not depicted in the table had no rooted cuttings. Statistical significance was determined at p < 0.05. A total of 540 stem cuttings were planted in each season. CJ: coconut juice.
| Sampling Season | Total Values | |||||
|---|---|---|---|---|---|---|
| Dependent Variables | Effects | Winter | Spring | Summer | Fall | |
| Rooting (%) | CJ | - | <0.0001 | 0.943 | <0.0001 | <0.0001 |
| Season | - | - | - | - | <0.0001 | |
| Substrate | - | 0.068 | 0.765 | 0.086 | 0.0300 | |
| CJ × Season | - | - | - | - | <0.0001 | |
| CJ × Substrate | - | 0.240 | 0.765 | 0.978 | 0.0483 | |
| Season × Substrate | - | - | - | - | 0.520 | |
| CJ × Season × Substrate | 0.465 | |||||
| Essential oil (%) | CJ | <0.001 | 0.012 | - | 0.004 | <0.0001 |
| Season | - | - | - | - | 0.003 | |
| Substrate | 0.260 | 0.279 | - | 0.649 | 0.001 | |
| CJ × Season | - | - | - | - | 0.043 | |
| CJ × Substrate | 0.405 | 0.245 | - | 0.256 | 0.001 | |
| Season × Substrate | - | - | - | - | 0.279 | |
| CJ × Season × Substrate | 0.355 | |||||
Type and percentage of the individual components obtained from the EOs of the fresh samples in each season.
| Winter | Spring | Summer | Fall | ||||
|---|---|---|---|---|---|---|---|
| Type | % | Type | % | Type | % | Type | % |
| Sabinene | 31.22 | Nerolidol | 8.58 | 3-Carene | 16.87 | Beta-pinene | 34.76 |
| Citronellol | 12.59 | Delta-cadinol | 7.59 | Terpinen-4-ol | 9.27 | Terpinen-4-ol | 8.80 |
| Terpinen-4-ol | 6.96 | Nerolidol E | 5.71 | Carvacrol | 8.99 | Myrcene | 6.18 |
| Germacrene D-4-ol | 5.27 | Methyl Citronellate | 4.52 | Germacrene D-4-ol | 4.27 | Germacrene D-4-ol | 3.69 |
| Myrcene | 4.77 | Alpha-terpineol | 4.38 | Gamma-terpinene | 3.5 | Gamma-terpinene | 3.18 |
| Methyl Citronellate | 3.76 | Farnesol | 3.28 | Myrcene | 3.22 | Citronellol | 2.99 |
| Elemol | 2.29 | Cubenol | 3.15 | Citronellol | 2.99 | Alpha-bisabolol | 2.64 |
| Citronellal | 2.25 | 2:4 Decadien-1-ol | 2.9 | Limonene | 2.74 | Delta Cadinene | 2.44 |
| Gamma-terpinene | 2.08 | Alpha-cadinol | 2.39 | Elemol | 2.62 | Linalool | 2.36 |
| Alpha-pinene | 2.06 | Terpinolene | 1.93 | Delta-cadinene | 2.35 | Alpha-pinene | 2.20 |
| Delta Cadinene | 1.99 | Terpineol (cis-beta) | 1.75 | Delta Cadinene | 1.79 | Limonene | 2.13 |
| Limonene | 1.93 | Myrcene | 1.18 | Terpinene (alpha) | 1.78 | Elemol | 2.08 |
| Linalool | 1.84 | Cyclogeraniol alpha | 0.83 | Viridiflorol | 1.76 | Alpha-Terpinene | 1.95 |
| Alpha Thujene | 1.69 | Cineole | 0.75 | Terpinolene | 1.57 | Alpha-Thujene | 1.91 |
| Alpha Bisabolol | 1.47 | Gamma Muurolene | 0.62 | p-Cineole | 1.37 | Citronellyl formate | 1.75 |
| Terpinolene | 1.27 | Alpha Copaene | 0.58 | Alpha Cadinol | 1.29 | Teroinolene | 1.60 |
| Alpha Terpinene | 1.26 | Muurolene | 0.51 | Thujene (alpha) | 1.21 | Cis-beta-Terpineol | 1.29 |
| Alpha Cadinol | 0.66 | 4-Decen-1-ol | 0.50 | Linalool | 1.21 | Gamma-Decalactone | 1.29 |
| Cis-beta Terpineol | 0.58 | Citronellol | 0.47 | Caryophyllene oxide | 0.94 | Alpha Cadinol | 1.08 |
| Alpha Terpineol | 0.41 | Pinene alpha | 0.46 | Methyl eugenol | 0.92 | 2-undecanone | 0.89 |
| Beta cis Ocimene | 0.39 | Linalool acetate | 0.45 | Aromadendrene | 0.90 | Citronellal | 0.88 |
| Germacrene D | 0.36 | Aromadendrene | 0.43 | Cis-beta-Terpineol | 0.76 | E-beta-Ocimene | 0.82 |
| Beta pinene | 0.19 | Terpinene alpha | 0.4 | Cymene (para) | 0.72 | Alpha Muurolene | 0.75 |
| Ortho Cymene | 0.13 | Alpha Amorphen | 0.38 | Alpha Muurolene | 0.69 | Limonene oxide | 0.65 |
| Carvacrol | 0.13 | Gurjunene | 0.27 | Thymol acetate | 0.58 | Alpha-Terpineol | 0.63 |
| Beta Elemene | 0.12 | Gamma Eudesmol | 0.25 | Carene (delta-3) | 0.38 | Linalool acetate | 0.60 |
| Caryophyllene | 0.1 | Humulene | 0.24 | Alpha selinene | 0.34 | Dec-9-en-1-ol | 0.46 |
| Caryophyllene oxide | 0.1 | Methyl geranate | 0.23 | Alpha Himachalene | 0.33 | Methyl geranate | 0.31 |
| Alpha phellandrene | 0.09 | Elemol | 0.23 | Gamma Cadinene | 0.3 | Cubenol | 0.24 |
| Gamma Muurolene | 0.09 | Undecanol | 0.22 | Carotol | 0.25 | Delta-3-Carene | 0.23 |
| Citronellyl acetate | 0.08 | Gamma Elemene | 0.22 | Methyl Generate | 0.24 | trans-Piperitol | 0.22 |
| Alpha Fenchene | 0.06 | Ocimene | 0.20 | Ocimene | 0.22 | Geraniol | 0.17 |
| Nonanal | 0.06 | Eugenol | 0.19 | Cubenol | 0.21 | p-Cymene | 0.15 |
| Neryl acetate | 0.06 | Limonene | 0.17 | Beta Phellandrene | 0.17 | Beta-phellandrene | 0.13 |
| Camphene | 0.05 | Citronellal | 0.11 | Spathulenol | 0.17 | Ledol | 0.12 |
| Terpinyl acetate | 0.05 | Caryophyllene | 0.11 | Camphene | 0.12 | Terpineol acetate | 0.11 |
| Alpha Humulene | 0.05 | Terpinene gamma | 0.04 | Beta Elemene | 0.11 | Germacrene D | 0.09 |
| Isopulegol | 0.03 | Hexanal | 0.02 | Alpha Amorphene | 0.11 | Gamma Eudesmol | 0.09 |
| Hexanal | 0.02 | Germacrene B | 0.11 | Thujone | 0.09 | ||
| n-Octane | 0.01 | Nonanal | 0.08 | Nonanal | 0.08 | ||
| Gamma Muurolene | 0.08 | Alpha-Fenchene | 0.05 | ||||
| Germacrene D | 0.07 | Camohene | 0.05 | ||||
| Cis-3-Hexenol | 0.06 | Rose oxide | 0.04 | ||||
| Hexanal | 0.04 | Germacrene B | 0.03 | ||||
| Octanol | 0.02 | ||||||
| Hexanal | 0.02 | ||||||
| Octane | 0.01 | ||||||
| Hexanol | 0.01 | ||||||
| z-3-Hexenol | 0.01 | ||||||
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; Zarei, Hossein 3 ; Zahedi, Bahman 1 ; Abdolhossein Rezaei Nejad 1 1 Department of Horticulture Sciences, Faculty of Agriculture, Lorestan University, P.O. Box 46, Khorramabad 68151-44316, Iran;
2 Department of Agroforestry Technology and Science and Genetics, Higher Technical School of Agricultural and Forestry Engineering and Biotechnology, University of Castilla-La Mancha, 02071 Albacete, Spain
3 Department of Horticulture Sciences, Collage of Plant Production, Gorgan University of Agricultural Sciences and Natural Resources, Gorgan 49138-15739, Iran;