¹ Institute of Botany of Academy of Sciences of the Republic of Uzbekistan, Tashkent, 100125, Uzbekistan
² Department of Biology, Faculty of Biotechnology, Namangan State University, Namangan, Uzbekistan
³ Department of Botany, University of Otago, Dunedin, New Zealand
⁴ The Botanical garden of the Republic of Uzbekistan, Tashkent, 100125, Uzbekistan
⁵ Institute of Botany of Academy of Sciences of the Republic of Uzbekistan, Tashkent, 100125, Uzbekistan
⁶ Department of Microbiology, Virology and Immunology, Bukhara State Medical Institute, Bukhara, Uzbekistan
⁷ 6Jizzakh State pedagogical university, Jizzakh, Uzbekistan
⁸ Institute of Botany of Academy of Sciences of the Republic of Uzbekistan, Tashkent, 100125, Uzbekistan
⁹ Department of General Medical Sciences, Faculty of Medicine, Namangan State University, Namangan, Uzbekistan
¹⁰ Biotechnology Department, Faculty of Sciences, Niğde Ömer Halisdemir University, Nigde, Turkey
¹¹ Institute of Botany of Academy of Sciences of the Republic of Uzbekistan, Tashkent, 100125, Uzbekistan
¹² Institute of Botany of Academy of Sciences of the Republic of Uzbekistan, Tashkent, 100125, Uzbekistan

Dilafruz Jamalova
Roles: Conceptualization, Investigation, Methodology, Writing – Original Draft Preparation

Mohizar Mirzaolimova
Roles: Conceptualization, Data Curation, Investigation, Methodology, Resources

Amir Zarekarizi
Roles: Conceptualization, Formal Analysis, Software, Validation

Feruza Mustafina
Roles: Data Curation, Formal Analysis, Methodology, Supervision

Vasila Sharipova
Roles: Conceptualization, Formal Analysis, Validation

Khilola Ubaydullayeva
Roles: Conceptualization, Writing – Review & Editing

Mukhayyo Kholdorova
Roles: Data Curation, Formal Analysis, Investigation

Gulsauir Kurbaniyazova
Roles: Conceptualization, Investigation, Methodology

Mirzohid Mirzaolimov
Roles: Data Curation, Software, Validation

Huseyin Turker
Roles: Validation, Writing – Review & Editing

Ziyoviddin Yusupov
Roles: Project Administration, Supervision

Komiljon Tojibaev
Roles: Formal Analysis, Project Administration, Supervision

Mature seeds of D. microcarpum were collected in July 2022 from the Fergana Valley, Namangan, Davlatabad district, (Girvansai street) on Kokunbai road, (300–500 m above sea level, N 71 ^°599538; E 41 ^°01789). Seeds of the second species, D. sabulosum, were collected in July 2023 near the tugain-sandy reserve located in the middle reaches of the Amu Darya River, Bukhara region of the Republic of Uzbekistan (N 40°3402; E 62°1314). The collected seeds were stored in closed paper bags at −10 °C and later transferred to a laboratory refrigerator for cold stratification at +4 °C for one month before processing. Seeds were treated with 70% ethanol for 1.5 min, then rinsed with sterile water for three times and then surface sterilized in 4% sodium hypochlorid solution for 20 min, followed by three washes with sterile water. The sterilized seeds were cultured on ј MS (Murashige and Skoog, 1962) medium supplemented with 30 g/l sucrose and 8 g/l agar. To overcome dormancy, the seeds were kept in a refrigerator (+4 °C) for 60 days. After cold stratification, the seeds were incubated at +23 ± 2 ^° C under 16/8 h photoperiod provided by cool fluorescent lamps in culture room. The segments of the germinated seeds were used as a source of explants.

The followings were used as explants: mature zygotic embryos, segments of germinated seeds (roots, hypocotyl, and cotyledons); MS media with combinations of 2,4 D (0, 0.5, 1 mg/l), benzylaminopurine (0.2, 0.5, and 1 mg/l), indole 3-butyric acid (0.2, 0.5, and 1 mg/l); α-naphthaleneacetic acid (0.2, 0.5, and 1 mg/l); tidiazuron (0.2, 0.5, and 1 mg/l) and kinetin (0, 0.2, 0.5 or 1 mg/l) were tested. The pH of the medium was adjusted to 5.8, and growth regulators were added post-autoclaving. Germinated seeds (root, hypocotyl, and cotyledon segments) were cut by 0.5–1 cm in length. The explants were cultured in sterilized Petri dishes containing 25 ml of above-mentioned media combined with required doses of growth regulators and without them as a control.

Aseptically isolated mature zygotic embryos and the segments of the germinated seeds (root, hypocotyl, and cotyledon) were used as explants: root and hypocotyl were of 10 mm, cotyledon of 5 mm length. The 14–20 days-old seedlings were cut and the segments were cultivated on sterile MS medium with 2,4-D (0–0.5 mg/l), IBA (0–0.5 mg/l), in combination with TDZ (0.2, 0.5, and 1 mg/l) TDZ, Kin (0, 0.2, 0.5, and 1 mg/l), and BAP (0–1 mg/l) for callus induction. Calli were subcultured on the same medium every 4 weeks, each time divided into pieces of 2–4 mm diameter.

Mature zygotic embryos were placed on MS medium without a growth regulator to induce direct somatic embryogenesis. Cultures were incubated at +23 ± 2 ° C with 1,800 lux provided by cool fluorescent lamps with a photoperiod of 16/8 hours in culture room. Each experiment was performed with 5 replicates and the percentage of callus induction in the different media was recorded in eight weeks.

In two months after cultivation, the embryogenic callus was transferred to MS medium without growth regulators for induction of the embryogenesis. Mature somatic embryos were detected in 4 weeks after the incubation of the explants of nutrient media. The rate of indirect embryogenesis and mean number of embryos per explant was also calculated after 16 weeks. Then the mature somatic embryos were transferred to a hormone-free MS medium for germination and plantlet formation.

Morphogenic calli were transferred to the regeneration medium with various concentrations of BAP (0.1–1 mg/l), or combined with IBA (0–0.5 mg/l) or NAA (0–0.5 mg/l), to evaluate plants regeneration.

The share of explants with embryogenic callus was recorded in 4 weeks after culture. At this stage, the share of callogenesis, indirect somatic embryogenesis and number of embryos at different stages (globular, torpedo and cotyledonary), conversion of embryos into seedlings were also recorded.

Data analysis was displayed with box plots to reflect the distribution, mean, and standard deviation of measurements, conducted in triplicate (n = 3). Statistical analysis was done with one-way analysis of variance (ANOVA), followed by Tukey’s multiple comparison post hoc test with a 95% significance level (P < 0.05) for datasets that met normality assumptions. For datasets demonstrating non-normal distribution, exponential transformation was applied. When normalization was not achievable, the Kruskal-Wallis test, accompanied by Dunn’s test and Benjamini-Hochberg (BH) procedure, was applied. All statistical analyses were performed using RStudio, version 2023.06.1.

The initial experimental approach aimed to use mature zygotic embryos as explants for both D. microcarpum and D. sabulosum to expedite the process by bypassing seed germination. However, the two species showed markedly different responses, demanding species-specific adaptations in explant selection. For D. microcarpum, zygotic embryos showed no response, failing to induce callus or plant formation. This lack of reactivity prompted a shift in strategy, leading to the use of seedling segments derived from germinated seeds as the primary explant source for this species. In contrast, zygotic embryos proved to be functional explants for D. sabulosum, and the lake of in vitro germination of D. sabulosum seeds validated the continued use of its zygotic embryos as explants. The decision to employ distinct explants for each species was driven by these differential responses.

Mature zygotic embryos and the segments of the germinated seeds of two species D. microcarpum and D. sabulosum were used to induce callusogenesis. The callus formation was observed on root explants of D. microcarpum in 7 days after in vitro cultivation (Fig. 2A), in 15 days on hypocotyl segments of the germinated seeds, and no callus was observed on cotyledon segments. Callusogenesis was observed on the the zygotic embryos of D. sabulosum within 8 to 10 days. On hypocotyl segments callus primarily formed at the basal ends, whereas on roots callus was detected across the entire the surface as well as on zygotic embryos ( Figure 2). Root-derived callus was white, watery with a smooth surface, gradually becoming brown, whereas hypocotyl-derived callus was soft or friable, exhibiting a sticky consistency and nodular form with proembryogenic structures, ranging from yellow-white to creamy color ( Figure 3: 3B).

The Kruskal-Wallis test results indicated significant differences in the impact of growth regulators on the callus induction in roots (χ² = 51.13, p = 0.007) and hypocotyls (χ² = 45.91, p = 0.024) in D. microcarpum. Further analysis using Dunn’s post-hoc test identified significant treatment effects in D. microcarpum compared to controls. For root explants, the application of 2,4-D (0.2 mg l⁻¹) combined with TDZ (0.2 mg l⁻¹) (93.3 ± 6.7%, Z = 3.47, p = 0.029) and combination of 2,4-D (0.5 mg l⁻¹) + KIN (0.2 mg l⁻¹) (93.3 ± 6.7%, Z = 3.47, p = 0.038) significantly increased callus induction over the control group ( Figure 3).

The Kruskal-Wallis test also revealed significant differences in callus induction across treatments in D. microcarpum hypocotyls (χ² = 45.91, p = 0.0240). However, subsequent pairwise comparisons using Dunn’s test did not identify any specific growth regulator combinations that significantly differed from each other after correcting for multiple comparisons (Fig. 4. B). However maximum callus induction (73.3 ± 6.7%) was observed in treatments 2,4-D (0.2 mg l⁻¹) + Kin (0.2 mg l⁻¹) (73.3 ± 6.7%), 2,4-D (1.0 mg l⁻¹) + Kin (0.2 mg l⁻¹) (73.3 ± 13.3%), 2,4-D (0.5 mg l⁻¹) + TDZ (0.2 mg l⁻¹) (73.3 ± 6.7%) and 2,4-D (0.5 mg l⁻¹) + TDZ (0.5 mg l⁻¹) (73.3 ± 6.7%). In contrast, no significant differences were detected in the callus induction of D. sabulosum across different treatments (χ² = 30.40, p = 0.3943). However, the maximum percentage of callus (93.3 ± 6.7%was observed in the treatment of 2,4-D (0.5 mg l⁻¹) + TDZ (0.5 mg l⁻¹) ( Figure 4).

The study demonstrated significant variations in somatic embryogenesis responses across different growth regulator combinations in D. microcarpum. The most effective treatment, 2,4-D (0.5 mg l⁻¹) combined with Kinetin (0.5 mg l⁻¹), achieved the highest mean percentage of somatic embryogenesis at 66.67 ± 6.67% and the greatest average number of somatic embryos at 7.67 ± 0.33 per explant. 2,4-D (0.2 mg l⁻¹) with Kin (0.5 mg l⁻¹) and 2,4-D (0.2 mg l⁻¹) with Kin (0.2 mg l⁻¹), yielding average embryo numbers of 3.00 ± 1.00) and 3.33 ± 0.33) respectively ( Figure 5).

Statistical analysis using the Kruskal-Wallis test confirmed significant differences both in embryogenic callus percentages (χ² = 57.01, p = 0.0014) and somatic embryo numbers (χІ = 59.13, p < 0.0008) across treatments. Dunn’s post-hoc test further highlighted the superiority of 2,4-D (0.5 mg l⁻¹) + Kin (0.5 mg l⁻¹) over less effective treatments like 2,4-D (0.2 mg l⁻¹) + KIN (0 mg l⁻¹) and the control, with significant pairwise differences.

Upon transferring embryogenic calluses to hormone-free media in 10 weeks, microscopic examination revealed somatic embryos at various developmental stages, including globular, heart-shaped, torpedo, and cotyledonary forms. While most embryos exhibited normal morphology, some abnormalities occurred, such as embryos with multiple cotyledons, and sporadic secondary embryos developing on primary embryos.

D. sabulosum did not produce embryogenic calluses under any phytohormone combinations tested. Among the species analyzed, only D. microcarpum achieved a frequency of somatic embryogenesis ranging from 5% to 40%. After six weeks on a hormone-free medium, approximately 40–50% of cotyledonary somatic embryos germinated and developed into phenotypically consistent young plantlets, showcasing the potential for efficient clonal propagation and subsequent transplantation into pots for further growth and analysis ( Figure 2. C, D).

Following the production of primary somatic embryos, the embryogenic callus of D. microcarpum was transferred to MS medium supplemented with various concentrations of growth hormones to induce secondary somatic embryogenesis. One-way ANOVA revealed significant differences in the mean number of secondary somatic embryos among the different growth regulator treatments (p = 0.02). The control treatment produces the highest average number of secondary somatic embryos (15.33 ± 2.60) (Fig. 6). This was followed by 2,4-D (0.2 mg l⁻¹) + Kin (0.5 mg l⁻¹) and IBA (0.2 mg l⁻¹) + BAP (0.5 mg l⁻¹), both producing means of 11.67 ± 0.88 and 11.67 ± 1.45 embryos, respectively. In contrast, NAA (0.2 mg l⁻¹) + BAP (0.5 mg l⁻¹) resulted in the lowest production of secondary embryos (6.33 ± 0.33), significantly differing from the control ( Figure 6).

The study evaluated the effectiveness of various growth regulator combinations on shoot and root formation in callus cultures of D. sabulosum using MS medium supplemented with different concentrations of BAP, NAA, and IBA. Calli obtained from zygotic embryos were quickly responsive, expanding and developing green spots within two weeks, which turned brown by the third week (Fig. 3 A and B). By the fifth week, green shoots began regenerating, with regeneration outcomes closely tied to the growth regulators used ( Figure 3. E and F).

The optimal shoot regeneration was observed with the combination of IBA (0.5 mg l⁻¹) and BAP (0.5 mg l⁻¹), achieving a regeneration rate of 66.7 ± 6.7% (Fig. 7. A). This was followed by IBA (0.2 mg l⁻¹) with BAP (0.2 mg l⁻¹), which resulted in a 53.3 ± 6.7% regeneration rate. Among BAP-only treatments, BAP (0.5 mg l⁻¹) yielded 40 ± 11.6% shoot regeneration, while the control group showed a modest regeneration rate of 26.67 ± 6.67% ( Figure 7).

The Kruskal-Wallis test for shoot regeneration did not show significant overall differences among treatments (χ² = 19.06, p = 0.060), but Dunn’s test revealed significant pairwise differences, notably between BAP (1.0 mg l⁻¹) and IBA (0.5 mg l⁻¹) + BAP (0.5 mg l⁻¹) (Z = −3.06, p = 0.037), underscoring the influence of specific hormone combinations on regeneration outcomes.

For root formation, significant differences in the number of roots per shoot were observed (χ² = 25.18, p = 0.0086). Control shoots produced an average of 1.3 ± 0.3 roots per shoot. Different concentrations of BAP alone (0.2, 0.5, and 1.0 mg l⁻¹) inhibited root formation. This stark contrast with the control was statistically significant for all BAP concentrations (Dunn’s test, p < 0.05).

Combinations of hormones generally promote root formation. The IBA (0.5 mg l⁻¹) + BAP (0.2 mg l⁻¹) treatment yielded the highest mean number of roots per shoot (1.7 ± 0.3). Other IBA and BAP combinations, including IBA (0.2 mg l⁻¹) + BAP (0.2 mg l⁻¹), IBA (0.5 mg l⁻¹) + BAP (0.5 mg l⁻¹), and IBA (0.2 mg l⁻¹) + BAP (0.5 mg l⁻¹), all produced means equal to the control (1.3 ± 0.3 roots per shoot) ( Figure 7B). Post-hoc analysis revealed significant differences between BAP-only treatments and several hormone combinations. The most pronounced difference was observed between BAP (0.2) and IBA (0.5) + BAP (0.2) treatments (Z = −2.869428, p = 0.04523361). This comparison underscores the contrasting effects of BAP alone versus its combination with IBA on root formation.

All data generated or analyzed during this study are included in this published article.

This research was supported by the the projects “Digital Nature. Development of a digital platform for the flora of Central Uzbekistan,” implemented by the Institute of Botany of the Academy of Sciences of the Republic of Uzbekistan for the period 2025–2029, and the project Development of the Digital Platform of the Flora of the Ferghana Valley as a ‘Nature Imperilled Area’ (Project No. AL-9224104319). This research was also supported by the project titled “Assessing climate change adaptation in endangered plants of Uzbekistan: A DNA barcoding approach” (AL 9224104464).

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