Gloriosa superba L., commonly known as glory lily and belonging to the Liliaceae family, holds significant importance as both a medicinal and ornamental plant (Gurung et al. 2021; Pallavi et al. 2022). Widely distributed across tropical and sub-tropical regions in Africa and Southeast Asia, this botanical marvel serves as a rich repository of specialized metabolites, prominently featuring colchicine—a high-value alkaloid (Ade and Rai 2009; Mosoh et al. 2024a; Umavathi et al. 2020). The distribution of colchicine varies across plant parts, with the seeds and tubers exhibiting the highest content, a trait influenced by geographical locations and species diversity (Mosoh et al. 2023). The plant’s traditional applications in Indian medicine span a broad spectrum, encompassing treatments for ailments such as gout, rheumatic arthritis, snake and insect bites, intermittent fevers, and various other medical conditions (Mosoh et al. 2024a).

However, the escalating demand for Gloriosa superba L., driven by the commercial interest in colchicine production, has led to its depletion in the wild, exacerbated by challenges like low seed set, dual seed dormancy, and limited tuber germination rates (Arumugam and Gopinath 2012; Jana and Shekhawat 2011; Mahajan et al. 2016; Mosoh et al. 2023; Padmapriya et al. 2016; Roy 2017; Samy et al. 2008; Sivakumar et al. 2003; Sivakumar and Krishnamurthy 2004; Somani et al. 1989; Swapna and Nikhila 2018; Yadav et al. 2016).

Plant tissue culture techniques can provide a new alternative to conventional methods by enabling in vitro clonal propagation. Conventional breeding methods have drawbacks such as high labor and time requirements and inefficiency (Mosoh et al. 2024c; Nadalizadeh Ghannad et al. 2023). Plant tissue culture has great potential for more efficient propagation and improvement, but there is no universally accepted regeneration method, so customized protocols are necessary (Kolar and Ghouse 2014; Sivakumar and Krishnamurthy 2000; 2004; Swapna and Nikhila 2018).

The utilization of in vitro plant production protocols, which involve both direct and indirect morphogenesis techniques, exhibits significant promise in the context of medicinal species such as Gloriosa superba L. (Dey et al. 2022). This methodology facilitates the extensive clonal multiplication of better genotypes and the enhancement of genotypes through mutagenesis and genetic engineering techniques (Nadalizadeh et al. 2023). A plant regeneration system that is both stable and efficient serves multiple purposes, including the ability to micropropagate and conserve endangered Gloriosa cultivars, as well as enabling the introduction of new features through genetic transformation (Li et al. 2023).

Although previous in vitro studies on Gloriosa superba L. have been conducted, the existing literature predominantly emphasizes the induction of calli in various plant parts such as the tubers and corms, revealing distinct induction rates and susceptibilities to browning (Akter et al. 2014; Anandhi and Rajamani 2012a; 2012b; Anandhi et al. 2016; Arumugam and Gopinath 2012; Chatterjee and Ghosh 2015; Gopinath and Arumugam 2012; Gopinath et al. 2014; Hassan and Roy 2005; Jawahar et al. 2018; Kolar and Ghouse 2014; Kumar et al. 2015; Mahajan et al. 2016; Mosoh et al. 2023; Mosoh et al. 2024b; Muruganandam et al. 2019; Sanyal et al. 2022; Somani et al. 1989; Yadav et al. 2016). Sivakumar and Krishnamurthy (2004) previously demonstrated successful indirect organogenesis using Gloriosa superba L. leaf explants, employing benzyl adenine (BA) or 6-(γ, γ-dimethylallylamino) purine (2iP) in combination with adenine sulfate (ADS) and sodium citrate. However, their study did not provide data on the acclimatization process following organogenesis.

Moreover, the regeneration efficiency of calli is genotype and explant-type dependent, making universal applicability challenging (Sanyal et al. 2022). To the best of our knowledge, a significant gap persists in the research landscape regarding callus induction, specifically from leaf explants, indirect organogenesis arising from leaf callus, and acclimatization in Gloriosa superba L. Limited studies on indirect organogenesis using Gloriosa superba L. leaf explants have been conducted thus far, and those that exist report slow regeneration efficiency (Sivakumar and Krishnamurthy 2004). Closing this gap is crucial for advancing our understanding of tissue culture techniques in Gloriosa superba L., potentially unlocking new avenues for improving regeneration efficiency and facilitating the practical application of these techniques in plant propagation and conservation efforts.

This study focuses on developing an efficient protocol for the regeneration of Gloriosa superba L. using in vitro leaf callus culture (indirect organogenesis). The transplantation of these resulting plantlets into natural environments is crucial to filling critical knowledge gaps. The outcome of this study not only helps to conserve this endangered species but also lays the groundwork for further optimization and the introduction of new traits through genetic transformation. The study is a significant contribution to the fields of medicinal plant research, biodiversity conservation, and sustainable agriculture.

Plant materials and reagents

Gloriosa superba L. seeds were systematically harvested from robust and healthy mature plants within the confines of India’s Pachmarhi Biosphere Reserve (Fig. 1). The necessary plant growth regulators, including 6-Benzylaminopurine (BAP), Indole-3-acetic acid (IAA), Kinetin (KN), Gibberellic acid (GA₃), 1-Naphthalene acetic acid (NAA), Indole-3-butyric acid (IBA), Casein Hydrolysate (CH), and Coconut Water (CW), and requisite solvents, were procured from Sigma-Aldrich (Mumbai, India).

Figure 1. Photograph of glory lily (Gloriosa superba L.) in its natural habitat at the Pachmarhi Biosphere Reserve, Madhya Pradesh, India

Seed surface sterilization

The seeds underwent rigorous surface sterilization procedures to ensure optimal cleanliness and eliminate external contaminants. Initially, they were thoroughly cleansed under running tap water for 15 minutes. Subsequently, the explants (seeds) underwent an 8-minute wash with a 5% (v/v) Teepol solution, followed by five rinses with double-distilled water (DDW). Further sterilization involved treating the explants with a 2% (w/v) Bavistin solution for 10 minutes, followed by five additional rinses with DDW. The final stage of surface sterilization occurred within a laminar airflow chamber, where the explants (seeds) were exposed to 70% (v/v) ethanol for 20 seconds, followed by an 8-minute treatment with 0.15% (w/v) HgCl₂. Post-treatment, the explants underwent five thorough washes with sterile DDW to ensure the removal of any residual sterilizing agents.

In vitro seed germination

Following the incorporation of all medium constituents, excluding agar, the pH of the medium was meticulously adjusted to 5.8 using either 1N HCl or 1N NaOH. Subsequently, 0.8% agar (Himedia, Mumbai) was added, and culture flasks containing 50 ml of non-solid basal MS medium were hermetically sealed with non-absorbent cotton plugs before autoclaving for 20 minutes at 121 °C under 104 kPa pressure. Surface-sterile seeds were then placed with precision on half-strength MS medium supplemented with 1.5 mg L^-1 GA₃, 1.5 mg L^-1 BAP, and 3% (w/v) sucrose (Himedia, Mumbai). These seeds were cultured under precisely controlled conditions, maintaining a temperature of 25 ± 2 °C, a 16-hour photoperiod with an irradiance of 80 µmol m^-2 s^-1, and a relative humidity of 70%. Sterile leaf explants for the initiation of leaf callus were obtained from seedlings aged between 45 and 60 days.

Leaf callus induction

Leaf explants obtained from in vitro germinated seedlings were carefully washed with sterilized double-distilled water (DDW) for good measure, and subsequently trimmed into uniform sizes of 1cm by 1cm. The prepared explants were then placed under aseptic conditions onto full-strength Murashige and Skoog (MS) medium containing 3% (w/v) sucrose (HiMedia, Mumbai), 10 mg L^-1 casein hydrolysate (CH), 0.8% (w/v) agar (HiMedia, Mumbai) and pH 5.8. For each treatment group, different concentrations and combinations of BAP (0.5 and 1.0 mg L^-1), NAA (0.5, 1.0, 1.5, 2.0, 2.5, and 3.0 mg L^-1), and KN (0.5 and 1.0 mg L^-1) were added to the MS medium. Cultures were incubated in complete darkness at 25 ± 2°C and 60% relative humidity in a controlled chamber. After an incubation period of four weeks, several parameters were recorded in detail for further analysis. These parameters included the frequency and response rate of callus development from leaf explants, the time required for callus initiation, and the fresh and dry weight of the calli.

Induction of secondary differentiation of morphogenic callus cells

Upon successful callus induction, selected calli were randomly assigned and subjected to various treatments for shoot regeneration. The calli were cultured on freshly prepared, contamination-free MS media, consisting of 5 mg L^-1 CH, 20% CW (v/v), 0.8% agar (Himedia, India), and adjusted to pH 5.8. The media were supplemented with distinct combinations and concentrations of BAP (1.0, 1.5, 2.0, and 2.5 mg L^-1), KN (0.5 and 1.5 mg L^-1), and NAA (0.5 and 1.5 mg L^-1). These sterile cultures (leaf calli) were then incubated under controlled conditions with a light intensity of 20 µmol m^-2 s^-1, a photoperiod of 16/8 hour (day/night), and a temperature of 25 ± 2°C. After a four-week incubation period, meticulous assessments were conducted, encompassing parameters such as the frequency of shoot primordia formation and response rate (%), the number of shoot primordia and shoots per explant, as well as the fresh and dried weights of the resultant shoots.

Root induction

In vitro regenerated shoots of Gloriosa superba L. measuring 3-4 cm in height were meticulously excised and allocated to distinct root induction treatment groups. These shoots were subsequently immersed in freshly prepared sterile half-strength MS media supplemented with varying concentrations of IBA (0.5, 1.0, and 1.5 mg L^-1), IAA (0.5, 1.0, and 1.5 mg L^-1), and NAA (0.5, 1.0, and 1.5 mg L^-1). The control treatment comprised PGR-free, half-strength MS media. The rooting process took place under controlled conditions, with temperatures maintained at 25 ± 2°C, a photoperiod of 16/8 hours (light/dark), and an irradiance of 80 µmol m^-2 s^-1, provided by cool-day fluorescent lamps (TL 40W/54). Relative humidity levels were maintained within the range of 55 to 60%. Following a 4-week incubation period, various parameters such as root-to-shoot ratio, fresh weight, and dry weight were meticulously recorded.

Biomass determination

For fresh weight (mg) determination, harvested calli, shoots, and roots (after removing the base of the shoots) were weighed on a precision balance. To determine dry weight (mg), the same plant sample types were oven-dried at 45°C for 24 hours before being weighed again.

Acclimatization

Following successful root induction and development, the plantlets underwent a meticulous cleansing process, being delicately removed from the rooting media and thoroughly washed with deionized water to eliminate any residual media. Subsequently, the cleansed plantlets were carefully transplanted into petite containers with a 7 cm diameter, including options such as polyethylene bags, plastic trays, plastic pots, or thermocol cups. These containers were filled in a 1:1 ratio with a mixture of sterilized vermiculite and soil. To facilitate the initial acclimatization stage, the plantlets were subjected to a 16-hour photoperiod illuminated by white fluorescent tubes (40 W; Philips, Mumbai), providing a photosynthetic photon flux density of 50 µmol m^-2 s^-1.

Plantlets were shielded with transparent polyethylene bags featuring microscopic air holes to safeguard against dehydration and uphold optimal relative humidity (RH) levels. Within the culture room (CR), the temperature was meticulously maintained at 25 ± 2°C throughout the acclimatization process. To facilitate air exchange, the polyethylene bags were intermittently removed for one hour daily. For two weeks, potted plantlets received watering every four days with 10 ml of a half-strength MS basal salt solution devoid of sucrose or myo-inositol. During the subsequent stage of acclimatization (third to sixth week), the plantlets were transplanted into medium-sized containers—such as polyethylene bags, plastic vessels, or thermocol vessels—filled with a 2:1:1 ratio (v/v) mixture of garden soil, sand, and vermiculite. Subsequently, for a month-long duration, the plantlets were housed within a shade net house (SNH), where they received daily misting with tap water. During this phase, the RH was gradually reduced by half, ensuring a gradual adjustment to ambient environmental conditions.

Plant survival, along with various growth parameters including plant height (measured in centimetres), number of leaves per plant, number of flowers per plant, and number of microtubers per plant, were assessed two weeks post-transplantation into a sterilized substrate comprising vermiculite and soil in a 1:1 ratio within the culture room (CR). Additionally, evaluations of the same parameters were conducted one week after (during the third week) transplantation into garden soil, sand, and vermiculite mixture (2:1:1) under shade in the net house (USNH). Subsequently, the plantlets were transferred to larger earthen pots with a diameter of 15 cm, filled with a mixture of ordinary garden soil, sand, and farmyard manure in a 2:1:1 (v/v) ratio. For five weeks (until the eighth week), these plantlets were exposed to direct sunlight (DSL) to monitor their continued growth and development, and all the above-mentioned parameters were equally assessed and recorded.

Data collection spanned eight weeks after the commencement of plantlet acclimatization. During this period, weekly observations were conducted following the implantation of plantlets into different potting mixes, with four replicates employed for robust analysis. The survival percentage of the regenerated plantlets was determined using the formula: survival rate (%) = (Number of Surviving Regenerated Plants / Total Number of Transplanted Regenerated Plants) * 100%. All data presented in this study are expressed as mean values accompanied by their respective standard errors (SE).

Statistical analysis

A completely randomized experimental design was implemented. For instance, in the case of callus induction, 1 cm-long leaf explants were randomly allocated to treatment groups. Similarly, callus units and microshoots were randomly assigned to shooting and rooting treatment groups, respectively. Each treatment level was replicated four times, with eight leaf explants (callus induction), six callus units (shooting), or twelve microshoots per replicate (rooting), depending on the specific experiment. The trials were conducted twice to ensure the reliability and replicability of the results. Following four weeks of growth and development, data for all parameters were collected (Fig. 2, 4, 6).

Figure 2. Effects of concentrations of auxin (NAA) in combination with cytokinins (BAP and KN) on callus induction, growth, and subsequent biomass increase of Gloriosa superba L. leaf explants in Murashige and Skoog medium supplemented with 10 mg L^-1 casein hydrolysate. (A) Mean number of leaf explants that formed a callus. (B) Rate of callus induction. (C) Mean number of days required for callus induction. (D) Mean fresh weight. (E) Mean dry weight. Treatments are T1: 0.5 mg L^-1 NAA and 0.5 mg L^-1 BAP; T2: 1.0 mg L^-1 NAA and 0.5 mg L^-1 BAP; T3: 1.5 mg L^-1 NAA and 0.5 mg L^-1 BAP; T4: 2.0 mg L^-1 NAA and 1.0 mg L^-1 BAP; T5: 2.5 mg L^-1 NAA and 1.0 mg L^-1 BAP; T6: 3.0 mg L^-1 NAA and 1.0 mg L^-1 BAP; T7: 0.5 mg L^-1 NAA and 0.5 mg L^-1 KN; T8: 1.0 mg L^-1 NAA and 0.5 mg L^-1 KN; T9: 1.5 mg L^-1 NAA and 0.5 mg L^-1 KN; T10: 2.0 mg L^-1 NAA and 1.0 mg L^-1 KN; T11: 2.5 mg L^-1 NAA and 1.0 mg L^-1 KN; T12: 3.0 mg L^-1 NAA and 1.0 mg L^-1 KN. Bars indicate mean ± standard error. Different letters indicate significant differences determined via Tukey’s test at p ≤ 0.05. NAA, 1-naphthalene acetic acid; BAP, 6-benzylaminopurine; KN, kinetin

Figure 4. Effects of various PGR combinations on shoot regeneration via callus cultures generated from Gloriosa superba L. leaf explants in Murashige and Skoog medium supplemented with 5 mg L^-1 casein hydrolysate + 20% coconut water (v/v). (A) Mean number of calli that formed shoot primordia. (B) Rate of shoot primordium formation. (C) Mean number of shoot primordia per callus. (D) Mean number of shoots per callus. (E) Mean fresh weight. (F) Mean dry weight. Treatments are T1: 1.0 mg L^-1 BAP and 0.5 mg L^-1 KN; T2: 1.5 mg L^-1 BAP and 0.5 mg L^-1 KN; T3: 2.0 mg L^-1 BAP and 0.5 mg L^-1 KN; T4: 2.5 mg L^-1 BAP and 1.5 mg L^-1 KN; T5: 1.0 mg L^-1 BAP and 0.5 mg L^-1 NAA; T6: 1.5 mg L^-1 BAP and 0.5 mg L^-1 NAA; T7: 2.0 mg L^-1 BAP and 0.5 mg L^-1 NAA; T8: 2.5 mg L^-1 BAP and 1.5 mg L^-1 NAA; T9: control. Bars indicate mean ± standard error. Different letters indicate significant differences determined via Tukey’s test at p ≤ 0.05. PGR, plant growth regulator; NAA, 1-naphthalene acetic acid; BAP, 6-benzylaminopurine; KN, kinetin; control, culture medium without PGRs

Figure 6. Effects of different concentrations of auxins on in vitro rooting of Gloriosa superba L. microshoots in half-strength Murashige and Skoog medium. (A) Mean root-to-shoot ratio. (B) Mean fresh weight. (C) Mean dry weight. Treatments are T1: 0.5 mg L^-1 IBA; T2: 1.0 mg L^-1 IBA; T3: 1.5 mg L^-1 IBA; T4: 0.5 mg L^-1 IAA; T5: 1.0 mg L^-1 IAA; T6: 1.5 mg L^-1 IAA; T7: 0.5 mg L^-1 NAA; T8: 1.0 mg L^-1 NAA; T9: 1.5 mg L^-1 NAA; T10: control. Bars indicate mean ± standard error. Different letters indicate significant differences determined via Tukey’s test at p ≤ 0.05. IAA, indole 3-acetic acid; IBA, indole-3-butyric acid; NAA, 1-naphthalene acetic acid; control, culture medium without any plant growth regulators

The callus induction rate was assessed by dividing the number of leaf explants forming callus by the total number of replicates, then multiplying by 100 to obtain a percentage. Likewise, the rate of shoot primordia formation was determined by dividing the total number of shoot primordia-forming calli by the total number of replicates, also multiplied by 100 to express it as a percentage. Similarly, the root treatment response rate was calculated as the percentage of microshoots that successfully formed roots, obtained by dividing the total number of rooted microshoots by the total number of replicates and multiplying by 100.

The normality of the data was assessed using the Shapiro-Wilk test. Parametric tests, specifically one-way analysis of variance (ANOVA) at a significance level of α = 0.05, were applied when the normality test yielded a non-significant result (p ≥ 0.05), indicating normally distributed data. Conversely, non-parametric tests, such as the Kruskal-Wallis test at α = 0.05, were utilized when the normality test resulted in a significant outcome (p ≤ 0.05), indicating non-normally distributed data. Data analysis was performed using R Studio (version 4.4.0). Post-hoc mean separation to identify significant differences between treatment groups was achieved through Tukey’s honestly significant difference (HSD) test at α = 0.05. Mean values ± standard error were presented for all data (Fig. 2, 4, 6, 8). In the figures, different letters denote statistically significant differences at p ≤ 0.05, facilitating straightforward data interpretation and comparison.

Figure 8. Acclimatization of in vitro-regenerated Gloriosa superba L. microshoots. (A) Plant survival. (B) Plant height (cm). (C) Number of leaves per plant. (D) Number of microtubers per plant measured after two weeks of transplantation in sterilized substrate (vermiculite + soil, 1:1) in a CR, one week (i.e., on the third week) of transplantation in garden soil + sand + vermiculite (2:1:1) USNH, and five weeks (i.e., eighth week) of transplantation in garden soil + sand + vermiculite (2:1:1) under DSL. Bars indicate mean ± standard error. Different letters indicate significant differences determined via Tukey’s test at p ≤ 0.05. CR, culture room; USNH, under shade in net house; DSL, direct sunlight

This study investigates the effects of various plant growth regulator (PGR) combinations, specifically involving BAP, NAA, and KN on callogenesis (Fig. 2). The findings reveal a notable spectrum of variability in the initiation period for callus formation, callus induction rate (%), and callus weight (mg), owing to the diverse concentrations of auxins and cytokinins employed in the experimental setups. Additionally, the investigation extends to elucidating the role of PGRs in indirect organogenesis through callus formation (Fig. 4) and in vitro rooting (Fig. 6). Moreover, the research encompasses a thorough exploration of the acclimatization process (Fig. 8).

Callus formation, texture, and color

After a four-week culture cycle, comprehensive data was gathered. The results of the experiment revealed that the interactions of different combinations of PGRs significantly influenced key parameters, including callus induction rate, days required for callus initiation, and callus weight (Fig. 2A-2E).

In vitro, adding cytokinins caused a significant decrease in cell wall lignification, which helped the start of callus growth and its subsequent expansion. Typically, callus growth initiates at the incision site of the explant, progressively extending to envelop the entire explant surface (Fig. 3C). The initial coloration of the calli spanned from colorless to a light green shade, later transitioning to a yellowish-green tint. Across all treatments, the texture of the calli remained consistently friable. Notably, certain calli exhibited hyperhydric exudates leading to necrosis, while others displayed embryogenic properties, promptly fragmenting into min pieces (Fig. 3).

Figure 3. (A) Plantlets germinated in vitro from seeds. (B) Inoculation of dissected leaf explants onto callus induction medium. (C) Callus formation and proliferation from leaf explants. (D) Necrosis. Scale bar = 1 cm

Effects of PGRs on callus induction

The time of callus initiation, percentage of callus development, and fresh and dry weight of the calli varied between PGR combinations, with values ranging from 13 to 24.20 days, 34.38% to 71.88%, 144 to 300 mg, and 19.8% to 32.60%, respectively. MS medium supplemented with 1.5 mg L^-1 NAA + 0.5 mg L^-1 KN had the shortest callus initiation duration (12.2 days), followed by 1.0 mg L^-1 NAA + 0.5 mg L^-1 KN (13 days), 2.0 mg L^-1 NAA + 1.0 mg L^-1 KN (14.8 days), and 1.5 mg L^-1 NAA + 0.5 mg L^-1 BAP (15.5 days). These changes, however, were not statistically significant (Fig. 2C) (p > 0.05).

MS medium supplemented with 1.5 mg L^-1 NAA + 0.5 mg L^-1 KN, 1.0 mg L^-1 NAA + 0.5 mg L^-1 KN, and 1.5 mg L^-1 NAA + 0.5 mg L^-1 BAP concentrations were shown to be efficacious for callus induction, with rates of 71.88%, 65.63%, and 62.50%, respectively. The differences, however, were not statistically significant (Fig. 2A, 2B) (p < 0.134). In terms of callus weight, the 1.5 mg L^-1 NAA + 0.5 mg L^-1 KN treatment produced the greatest average fresh weight (300 mg), demonstrating a significant difference (p < 0.001) (Fig. 2D). Similarly, this treatment had the greatest average dry weight (32.6 mg), followed by 1.5 mg L^-1 NAA + 0.5 mg L^-1 BAP (31.6 mg) and 2.0 mg L^-1 NAA + 1.0 mg L^-1 KN (29.4 mg).

On the other hand, treatments with 3.0 mg L^-1 NAA + 1.0 mg L^-1 BAP or 3.0 mg L^-1 NAA + 1.0 mg L^-1 KN, 2.5 mg L^-1 NAA + 1.0 mg L^-1 BAP, and 0.5 mg L^-1 NAA + 0.5 mg L^-1 BAP required more than twenty days for callus initiation (24.2, 21.2, 20.8, and 20.5 days) with callus induction rates of 37.5%, 43.75%, 40.63%, and 34.38%, respectively. For these treatments, the mean callus fresh weight was 144 mg, 240 mg, 219 mg, and 148 mg, respectively.

Effects of plant growth regulators on in vitro shoot proliferation from callus

To enhance shoot proliferation, calli recovered from Gloriosa superba L. leaf explants were extracted and subcultured on an MS medium supplemented with different combinations and concentrations of plant growth regulators (PGRs) such as BAP, KN, and NAA. Adding cytokinin and auxin to the nutritional medium improved shoot primordia development, shoot growth (including the number of shoots), and shoot weight (Fig. 5).

Figure 5. (A) and (B) Initiation of shoot primordium (after 15 d) and callus with primordium (after one week). (C)-(F) Shooting and maturation of shoots from callus (after four weeks). Scale bar = 1 cm

Shoot regeneration began 14-28 days after culturing in this experiment. At regular intervals, additional shoot parameters were collected (Fig. 5A, 5B). The findings showed that combining PGRs (BAP, KN, and NAA) was critical and important in shoot regeneration via callus formation. MS media supplemented with 2.0 mg L^-1 BAP + 0.5 mg L^-1 NAA produced the greatest rate of shoot primordia production from callus (79.17%), a significant difference from the control treatment (PGR-free) (p < 0.01). This was followed by 2.0 mg L^-1 BAP + 0.5 mg L^-1 KN (70.83%) and 1.5 mg L^-1 BAP + 0.5 mg L^-1 NAA (66.67%) (Fig. 4A, 4B).

Moreover, the highest average number of shoot primordia per callus (6.12) (p < 0.001) and shoots per callus (5.25) (p < 0.008) were recorded in MS media supplemented with 2.0 mg L^-1 BAP + 0.5 mg L^-1 NAA, which exhibited significant differences compared to the control treatment (Fig. 4C, 4D). Furthermore, the MS medium supplemented with 2.0 mg L^-1 BAP + 0.5 mg L^-1 NAA yielded the greatest average fresh weight (216 mg) (p < 0.001) and dry weight (32.1 mg) (p < 0.001) of microshoots, with significant differences (Fig. 4E, 4F).

Effects of plant growth regulators on in vitro rooting

The current study investigated the effects of various types and concentrations of auxins on root induction rate, average root-to-shoot ratio, and root biomass weight in the nutrient medium. Various amounts of IAA, NAA, and IBA supplied in MS media were used to test in vitro root induction of callus-regenerated shoots.

When the media were supplemented with 1.0 mg L^-1 IBA, the greatest average root-to-shoot ratio, fresh weight, and dry weight (5.75, 143 mg, and 22.2 mg, respectively) were detected, with a significant difference compared to the PGR-free control treatment (p < 0.001) (Fig. 7). This was followed by 1.0 mg L^-1 IAA (5.00, 140 mg, and 19.9 mg, respectively) supplementation (Fig. 6A, 6B, 6C).

Figure 7. Rooting of plantlets in vitro in half-strength Murashige and Skoog medium supplemented with 1.0 mg L^-1 indole-3- butyric acid. (A) Initiation of rooting. (B) Root elongation and proliferation. Scale bar = 4 cm

Ex vitro acclimatization

Plantlets produced from leaf explants showed great acclimatization potential and were quickly treated to hardening treatments. When the plantlets were moved ex-vitro onto a substrate of sterilized vermiculite and soil (1:1, v/v) and put in a culture chamber for fourteen days, they acclimatized completely.

They were then shaded for seven days on a substrate made of garden soil mixed with sand and vermiculite (2:1:1, v/v) (Fig. 8A, 9A, 9A-9D). However, after five weeks of development on the same substrate in direct sunlight, the survival rate dropped to 76% (Fig. 8A, 9E). Notably, all surviving plants displayed normal characteristics, including reaching standard plant height, developing multiple leaves, and producing microtubers. These results revealed a statistically significant difference (p < 0.001) (Fig. 8B-8D, 9E).

Figure 9. Acclimatization of in vitro-regenerated Gloriosa superba L. microshoots. (A) Microshoots were transplanted into sterilized substrate (vermiculite + soil, 1:1) and grown in a culture room. (B) Plantlets were covered with polyethylene castings containing tiny punctures. (C) Plantlets were transplanted into medium-sized polyethylene vessels that contained a mixture of garden soil + sand + vermiculite (2:1:1) under shade in a net house. (D and E) Plantlet microtuber development at the later stages of acclimatization after growing in earthen vessels containing a mixture of garden soil + sand + vermiculite (2:1:1) under direct sunlight. Scale bar = 2 cm (A–D) and 8 cm (E)

The selection of leaf explants was deliberate, leveraging their well-documented ability to foster uniform cell development and mitigate potential variances induced by the culture conditions, as evidenced by previous studies (Chen et al. 1999; Horsch et al. 1985; Liu et al. 2022). This methodological choice establishes a robust framework for a reliable technique, facilitating callus induction from Gloriosa superba L. leaf explants, subsequent indirect organogenesis, and successful ex vitro acclimatization.

Efficiency is a crucial factor in plant tissue culture. It involves optimizing processes, workflows, and resource utilization to achieve tasks and produce plants at a low cost with minimum time and effort while maximizing productivity and output. Therefore, the efficiency of various plant tissue culture protocols for regenerating different or the same plant species can vary significantly based on factors such as time, number of steps, resource allocation, and costs. The most efficient protocol is considered the most advantageous from both conservation and industrial perspectives. In modern plant tissue culture, prioritizing efficiency in protocol development is crucial. Efficiency should be regarded not merely as an option but as an essential scientific requirement. This entails streamlining procedures wherever possible, such as achieving shooting and rooting in a single step, without compromising the performance metrics of shoot and root numbers or the resources, time, and effort expended (Mosoh et al. 2023). Applying such a paradigm can potentially improve productivity and resource utilization, benefiting both conservation efforts and commercial applications. As a result, this study aimed to improve the efficiency of indirect organogenesis and the regeneration of plants from callus, specifically those derived from Gloriosa superba L. leaf explants.

As far as we know, Sivakumar and Krishnamurthy (2004) are the only researchers to have successfully reported indirect organogenesis using Gloriosa superba L. leaf explants. These researchers obtained young leaf explants from five-week- old seedlings. The process took about five weeks, with two to three 7-day subculture intervals, to form calluses. Using 2,4-D and kinetin (0.452-4.64 µM), a 98% response rate was achieved. It took about 35 days for shoot morphogenesis from the callus in shooting media that contained ADS and 2iP (2.72-9.84 µM). After that, it took another 35 days for the shoots to proliferate in shoot multiplication media that contained BA mixed with ADS and sodium citrate (2.72-13.32 µM). Finally, rooting took another two weeks. Overall, five-week-old seedlings were used as the main source of leaf explants, and the whole process, from callus induction to rooting, took 119 days (17 weeks), not counting the time needed for acclimatization.

MS media, supplemented with varying concentrations and combinations of BAP, NAA, KN, and 2,4-D, are commonly employed for callus induction. Different phytohormonal combinations have been observed to yield varying types and degrees of callus formation (Dar et al. 2021; Mosoh et al. 2024b). BAP is a synthetic cytokinin that enhances plant growth by stimulating cell division. It plays a vital role in tissue culture research, promoting the growth and multiplication of shoots, and is commonly employed in medicinal plant tissue culture (Hemant Sharma 2018). Kinetin, also known as 6-furfuryl-aminopurine, acts as a plant cytokinin growth regulator. When incorporated into plant tissue culture conditions, it demonstrates the capacity to induce callus formation and subsequent tissue regeneration. Plant tissue culture protocols frequently employ kinetin alongside auxins in media formulations like Murashige and Skoog (Bouhouche and Ksiksi 2007). NAA is an important auxin commonly used in plant tissue culture for the micropropagation of various species. Its effectiveness extends to influencing morphogenetic effects such as shoot proliferation, callus induction, and root induction (Ahmad et al. 2022). In our previous study, we found that using plant growth regulators, specifically NAA combined with kinetin and casein hydrolysate, was highly effective in promoting callus formation. Moreover, we observed that BAP combined with NAA, along with additives such as coconut water and casein hydrolysate, showed remarkable efficiency in inducing shoots in non-dormant corm explants of Gloriosa superba L. Furthermore, IBA has been established as the most effective plant growth regulator for rooting in Gloriosa superba L. (Mosoh et al. 2023; Sivakumar and Krishnamurthy 2004).

Effects of PGRs on callus induction

In this study, leaf explants consistently exhibited the formation of friable and rapidly developing calli, demonstrating remarkable resilience to variations in the type, concentration, or combination of plant growth regulators applied (Fig. 2, 3). The supplementation of the MS growth medium with coconut water (CW) and casein hydrolysate (CH) significantly enhanced the efficiency of callus production. Notably, CW, enriched with cytokinins, played a pivotal role in promoting cell division and rapid growth, while CH proved crucial for inducing callus, fostering shoot development, and facilitating multiplication—a phenomenon well-supported by previous studies (Al-Khayri 2011; Bai and Qu 2001; Sinha and Roy 2004; Sridhar and Aswath 2014; Srinivasan et al. 2021).

The findings of this study underscore the intricate interplay between NAA and KN in callus induction, demonstrating their superior efficacy compared to the combination of NAA and BAP. The MS medium supplemented with 1.5 mg L^-1 NAA, 0.5 mg L^-1 KN, and CH proved optimal for callus formation, achieving results within 12 days. This is significantly faster than the 35 days reported by Sivakumar and Krishnamurthy (2004). However, Sivakumar and Krishnamurthy (2004) observed a higher callus response rate in Gloriosa superba L. leaf explants (98% compared to 71.88%). This superior performance may be attributed to their use of 2,4-D, a potent plant growth regulator (PGR) known for its efficacy in callus induction across various plant species, as well as multiple subculturing intervals in fresh media. However, the choice of 2,4-D likely contributed to the longer duration required for callus formation (Fitch and Moore 1990). This hypothesis is supported by our previous study, which reported callus induction within 9.50 days using NAA and Kinetin with CH as an additive, albeit in non-dormant corm bud explants of Gloriosa superba L. (Mosoh et al. 2024b). Notably, the addition of CH exerted a significant influence, particularly in the MS medium supplemented with 1.5 mg L^-1 NAA and 0.5 mg L^-1 KN, elucidating the complex relationship between auxins and cytokinins. These findings are consistent with previous studies that have highlighted the effectiveness of this combination in promoting callus formation (Kaviani et al. 2013; Liu et al. 1997; Okazawa et al. 1967; Saleem et al. 2022). However, a prior study examining the impact of auxins and Kinetin on callus growth in Haworthia aristata and Haworthia setata revealed that the combination of IAA (a natural auxin) and KN yielded favorable outcomes for both species. In contrast, the interaction between NAA (a synthetic auxin) and KN resulted in suboptimal and inconsistent results (Ogihara and Tsunewaki 1978). It is imperative to acknowledge the diverse effects of plant hormones across different species. Dar et al. (2021) compared various plant growth regulators and found that, out of all the growth regulators used, NAA exhibited a superior response in callus formation. The findings of Dar et al. (2021) regarding Atropa acuminate leaf explants serve as a poignant illustration of this variability. Such variations underscore the intricate interplay between plant hormones and species-specific physiological processes (Dar et al. 2021; Gaspar et al. 1996). Dar et al. (2021) concluded that, regardless of the type of explants used, the ratio and type of auxins and cytokinins play a critical role in callus induction.

Effects of plant growth regulators on in vitro shoot proliferation from callus

The most effective combination for shoot regeneration was identified as 2.0 mg L^-1 BAP and 0.5 mg L^-1 NAA, with the addition of 5 mg L^-1 CH, and 20% coconut water (v/v). This combination induced shooting within two weeks forming 5.25 shoots per callus unit without the shoot multiplication step using shoot proliferation media. Sivakumar and Krishnamurthy (2004) reported that after 35 days, 7 to 12 shoots were formed using a shooting medium supplemented with ADS and 2iP, followed by an additional 35-day shoot multiplication phase utilizing BA, ADS, and sodium citrate. Adenine sulfate (ADS) has been shown to greatly increase the number of multiple shoot inductions compared to culture media that do not contain ADS. This was observed in Carissa carandas (L.), an important medicinal plant (Imran et al. 2012). Moreover, Naaz et al. (2014) reported that applying BA was the most effective cytokinin for inducing shoot buds and shoots in Syzygium cumini L. shoot tip explants. However, the proliferated shoots exhibited slower and stunted growth, along with leaf abscission and shoot tip necrosis (STN). The addition of adenine sulfate (ADS) to the optimal medium significantly mitigated leaf abscission and STN issues, resulting in the production of up to 14 shoots (Naaz et al. 2014). These findings suggest that while ADS improves the overall quantity of shoots, it does not improve the shoot growth rate. 2iP is a cytokinin used in plant tissue culture to promote shoot morphogenesis and growth. According to Jana et al. (2013), cultures supplemented with 2iP exhibited elongated shoot lengths compared to those using other cytokinins such as BA, Kinetin, and Thidiazuron (TDZ). They found that 2iP was the most effective for shoot multiplication among the four. The efficacy of cytokinin type and concentration in stimulating shoot induction and elongation can vary depending on the plant species, which may also apply to Sophora tonkinensis (Jana et al. 2013). This conclusion is supported by another study on Gardenia jasminoides Ellis, which compared the effects of BA and 2iP on shoot proliferation and elongation. In this study, BA was found to be more effective than 2iP for both shoot multiplication and elongation (Chuenboonngarm et al. 2001). Therefore, the specific role of 2iP, beyond its promotion of shooting, remains unclear, while it is evident that ADS facilitated shoot multiplication in the study by Sivakumar and Krishnamurthy (2004). Despite the combined use of 2iP and ADS for shoot multiplication, it still required five weeks to initiate shoots, which is 2.5 times slower than the shoot initiation results reported in our study.

Interestingly, biomass accumulation was observed even in the PGR-free control treatment, highlighting the substantial contribution of CH and CW to in vitro shoot growth. This finding is consistent with previous studies that have also emphasized the importance of these additives (Salehi et al. 2017; Yong et al. 2009). However, biomass accumulation in the control was lower compared to treatments that included PGRs, CH, and CW (Treatments 1 to 8 in Fig. 4E, 4F). This suggests a synergistic effect between PGRs, CH, and CW. In this study, the optimal shooting medium for the leaf-derived callus of Gloriosa superba L., which enhanced shoot development and biomass accumulation, contained 2.0 mg L^-1 BAP, 0.5 mg L^-1 NAA, 5 mg L^-1 CH, and 20% (v/v) coconut water (Fig. 4E). An earlier study highlighted the positive effects of BAP in combination with NAA, which were found to be optimal for inducing the regeneration of high-quality shoots from leaf-derived calli of Gerbera jamesonii Bolus (Bhatia et al. 2008). This observation is consistent with previous literature indicating that an elevated cytokinin-to-auxin ratio promotes optimal shoot morphogenesis (Mosoh et al. 2024b; Motte et al. 2014). The intricate interplay of selected phytohormones, along with the supportive functions of CH and CW in shoot development, underscores the complexity of the regulatory pathways involved. These findings offer invaluable insights into the underlying mechanisms governing in vitro shoot regeneration protocols, warranting further investigation and refinement.

Effects of plant growth regulators on in vitro rooting

A robust root system is crucial for successful plant propagation, especially under in vitro conditions (Krupa-Malkiewicz and Mglosiek 2016; Miri 2020). This study emphasizes the significant influence of exogenous auxins, particularly 1 mg L^-1 IBA, on key parameters such as the average root-to-shoot ratio as well as the fresh and dry weight of roots. These findings corroborate previous research, highlighting the efficacy of IBA as a potent auxin for in vitro rooting in Gloriosa superba L. (Mosoh et al. 2023; 2024d; Rafique et al. 2012; Srinivasan et al. 2021; Venkatachalam et al. 2012). The consistent results across multiple studies emphasize IBA’s reliability in promoting robust root development under in vitro conditions for Gloriosa superba L.

Ex vitro acclimatization

The acclimatization process revealed a notable disparity in the survival rates of plantlets under distinct environmental conditions, with a higher percentage observed in controlled settings such as the culture room and shade house. Conversely, exposure to direct sunlight outside the shaded housing led to a decline in the survival rate. This phenomenon may be attributed to an elevation in photosynthetically active radiation (PAR), potentially influenced by alterations in the vermiculite ratio, as suggested by previous studies (Hoang et al. 2020; Mosoh et al. 2024b; Teixeira da Silva et al. 2017). Remarkably, despite the variability in survival rates, no discernible changes in physical or growth characteristics were observed among the acclimatized plants, indicating their normal developmental progression.

This pioneering study unveils an efficient protocol for callus formation, indirect organogenesis, and acclimatization using leaf explants of Gloriosa superba L., marking a significant advancement in the field. The successful implementation of this approach is attributed to the synergistic effects of casein hydrolysate (CH) and coconut water (CW), optimizing both callus formation and shoot regeneration in this plant species. The most effective medium for callus induction, achieving results in approximately 12 days, was MS medium supplemented with 1.5 mg L^-1 NAA, 0.5 mg L^-1 KN, and 10 mg L^-1 CH. For shoot morphogenesis and biomass enhancement, a medium consisting of 2.0 mg L^-1 BAP, 0.5 mg L^-1 NAA, 5 mg L^-1 CH, and 20% (v/v) coconut water proved highly effective, successfully inducing shoot primordia within just 14 days. Additionally, this investigation identified 1.0 mg L^-1 IBA as the most efficient auxin for in vitro shoot rooting. Following eight weeks of acclimatization, the plantlets exhibited an impressive 76% survival rate, highlighting the robustness and viability of the developed protocol.

The established protocol exhibits the capacity for rapid large-scale production of genetically uniform Gloriosa superba L. plants, establishing a robust in vitro system suitable for genetic transformation using Agrobacterium-mediated techniques. While the present study underscores significant advancements, further investigations are warranted to explore additional factors influencing tissue culture responses and to evaluate potential variations in outcomes under diverse environmental conditions. Future research should investigate the potential of using adenine sulfate (ADS) in combination with optimal plant growth regulators (PGRs) and additives, such as casein hydrolysate (CH) and coconut water (CW), to determine if high shoot multiplication can be achieved and sustained within two weeks.

This research represents a significant advancement in harnessing the full potential of tissue culture techniques for plant propagation and broader biotechnological applications. The findings not only enhance our understanding of Gloriosa superba L. but also lay the groundwork for future research aimed at optimizing and expanding the utility of tissue culture in plant sciences and biotechnology.

The USDA National Institute of Food and Agriculture has provided support for this study, specifically under Hatch project 7001563. The corresponding author thanks Chief MOSOH Paul Tandong and Chieftess Ateyim Espe MOSOH Ostensia Nkeng of PINYIN (Santa, North-West Region, Cameroon) for their tremendous support. The corresponding author thanks Mr. Adamou Musa (TX, USA) and Mr. Tetu Acha Samuel (MD, USA) for their prompt and invaluable assistance with procurement appropriation. We want to express our gratitude to Dr. Rohit Sharma, Founder of the Centre for Biodiversity Exploration and Conservation (CBEC), for his valuable contributions during the initial stages of this project. Additionally, we express our sincere appreciation to the reviewer(s) for their diligent review of our work, and for their insightful comments and suggestions, which have significantly enriched the quality of our manuscript.