ARTICLE   Open Access    

Overcoming dual seed dormancy and enhancing in vitro seedling development of Gloriosa superba (L.) with a targeted sterilization approach and plant growth regulator synergy

More Information
  • Received: 29 May 2024
    Revised: 13 July 2024
    Accepted: 29 July 2024
    Published online: 12 September 2024
    Tropical Plants  3 Article number: e031 (2024)  |  Cite this article
  • Gloriosa superba L. is a critically endangered tropical plant with significant medicinal, cultural, and ornamental value. However, seed dormancy represents a significant barrier to natural regeneration in wild plant populations.

    This study found that sterilizing seeds with 0.15% mercuric chloride (HgCl2) for 8 minutes effectively eliminated contamination, ensuring 100% seedling survival.

    Pre-treatment with HgCl2 weakened the seed coat and induced chemical scarification, enhancing in vitro seed germination. Moreover, the imbibed Hg2+ ions likely induced the upregulation of MIPS genes in the seeds, which subsequently complemented the action of plant growth regulators (PGRs) in enhancing seed germination, seedling growth, and development.

    The combination of GA3 and BAP was most effective for in vitro seed germination. The combination of BAP and NAA was most effective for seedling development and elongation, while IBA was the best for rooting microshoots derived from seedlings.

  • The status of Gloriosa superba L. in the wild has been declining due to over-collection and habitat destruction. Intrinsic severe seed dormancy and low germination rates hinder the cultivation process. To conserve this plant, in vitro culture protocols have been developed to enhance seed germination and seedling growth. An effective sterilization method involving 0.15% mercuric chloride (HgCl2) for 8 min was found to eliminate contamination and yield a 100% survival rate, resulting in disinfested seeds and robust seedling growth. The most successful treatment consisted of Murashige and Skoog (MS) medium with 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP, along with 4% sucrose under a 16-h photoperiod, which achieved the highest average in vitro seed germination (9.25 out of 12 seed explants) and an impressive overall seedling survival rate of 77.08% after 30 d. Subsequent growth of two-week-old seedlings on MS medium with 1.5 mg·L−1 BAP and 1.0 mg·L−1 NAA supplemented with 30 g·L−1 sucrose and a 16-h photoperiod resulted in the greatest average seedling length (5.83 cm) and seedling root length (4.08 cm) after four weeks. Transferring the excised shoots of in vitro-grown seedlings to half-strength MS medium with 1.0 mg·L−1 IBA led to maximum root induction (84.37%) and further enhanced root development. These in vitro-grown plantlets were successfully acclimatized and transplanted under field conditions, with a 60% survival rate after 11 weeks.
    Graphical Abstract
  • The tea plant (Camellia sinensis) was first discovered and utilized in China, where its tender leaves were processed into tea. Tea has become the second most popular beverage after water worldwide. Tea contains tea polyphenols, amino acids, vitamins, lipopolysaccharides and other nutrients, as well as potassium, calcium, magnesium, iron, fluorine and other trace elements, which have antioxidant, lipid-lowering, hypoglycaemic, anti-caries, enhance the body's immunity and other physiological regulatory functions[1]. Among them, fluorine is an element widely found in the earth's crust, mainly in the form of fluoride in the environment, is one of the essential trace elements for human beings, and is vital to the growth and development of human bones and teeth[2]. When the human body takes in an appropriate amount of fluorine, it can effectively prevent the formation of dental caries, enhance the absorption of calcium, phosphorus and other elements of the human body. Excessive intake of fluorine however will lead to chronic cumulative poisoning, damaged bone tissue, affect the function of various tissues and organs in the body, and cause harm to health[3]. Tea plants can absorb and accumulate fluoride from air, water and soil, mainly concentrated in the leaves, most of the fluoride in the leaves can be released into the tea soup and be absorbed by the human body, so the fluoride content in tea is closely related to human health[4]. Generally speaking, green tea, black tea, white tea, oolong tea and yellow tea are made from the young buds and shoots of the tea plant, and their fluorine content is low. Some dark tea made from leaves with lower maturity has lower fluorine content. However, the dark tea made from leaves with higher maturity has a high fluorine content, and therefore poses a risk of excessive fluorine[5]. Long-term drinking of dark tea with excessive fluorine content is a cause of tea-drinking fluorosis[6].

    Dark tea, with its smooth taste and digestive benefits, became an indispensable drink in the lives of the Chinese herders, who were mainly meat eaters[7]. Dark tea has also gained popularity in the wider population because of its important health-promoting effects, such as prevention of cardiovascular and cerebrovascular diseases, lowering blood pressure, and promoting weight loss and fat reduction[8]. Long-term consumption of dark tea is likely to cause fluorosis for two reasons: 1) Chinese border ethnic minorities generally use the boiling method to brew dark tea, which increases the leaching rate of fluoride[9] and leads to high fluoride levels in the human body; 2) most of the fresh tea leaves utilized to make dark tea are older and more mature leaves, which contain higher levels of fluoride than younger leaves[10]. The mature leaves of tea plants accumulate a large amount of fluoride, but can grow normally without fluoride poisoning, indicating that tea plants are able to accumulate and tolerate fluoride. Due to the problem of tea-drinking fluorosis, the excessive accumulation of fluoride in tea plants has attracted widespread attention[10]. It is important to investigate the mechanisms related to fluoride absorption, transportation, enrichment, and tolerance in tea plants to develop effective and practical management and control programs to reduce the fluoride content in tea and ensure its safety. Recent research on defluorination measures for tea has included preliminary screening of tea germplasm resources, management measures during tea plant cultivation, processing technologies, and tea brewing methods. In this review, we summarize the results of studies on how fluoride moves from the environment into tea plants and the factors affecting this process, how fluoride is transported in tea plants, the mechanisms of fluoride tolerance in tea plants, and current measures to reduce the fluoride content in tea.

    Tea plants are fluoride-accumulators with the ability to absorb and accumulate fluoride from the surrounding environment. The fluoride content in tea plants is significantly higher than that of other plants under similar growth conditions[11]. After fluoride is absorbed by the roots of tea plants, it is transferred to the above-ground parts, and is also transferred from the leaves downward (Fig. 1), but not from the above-ground parts to the below-ground parts (Fig. 2). External factors such as atmospheric, soil, and water conditions around the tea plant and internal factors can affect the fluoride content in different plant tissues.

    Figure 1.  Fluoride absorption by tea plants. Tea plants absorb fluoride from the atmosphere, soil, and water. Fluoride in the atmosphere is absorbed through the stomata or cuticle of the leaf epidermis. Fluoride ions and fluoride–aluminum complexes in soil and water are absorbed by the roots.
    Figure 2.  Fluoride transportation in tea plants. Fluoride absorbed by the leaves is transferred to the leaf tips and edges, chelates with metal ions, and the complexes are deposited in the leaves. Fluoride can be transferred from old leaves to new tips. Fluoride from soil and water can form complexes with organic acids and aluminum, which are stored in the leaves of tea plants. Fluoride from soil and water can also be individually absorbed and transported for storage in the leaves.

    Fluoride occurs in many forms in nature. Fluoride in the atmosphere mainly exists in the form of hydrogen fluoride, and that in the soil mainly exists in three forms: insoluble, exchangeable, and water-soluble fluoride. Atmospheric fluoride is mainly absorbed through the stomata of tea plant leaves or the cuticle of the epidermis, and its concentration is relatively low[12]. Insoluble fluoride and exchangeable fluoride in the soil cannot be absorbed by tea plants. Water-soluble fluoride is the main form absorbed by the root system of tea plants[13]. Several studies have detected a significant positive correlation between the fluoride content in tea and the water-soluble fluoride content in soil[14,15]. Water-soluble fluoride mainly refers to the fluoride ion (F) or fluoride complexes in soil and water solutions, including free F and fluoride complexed with ions. The water-soluble state has the strongest activity and the highest biological availability, so it is conducive to the migration of fluoride in the environment[16].

    The root system is the main organ responsible for fluoride absorption in tea plants. Water-soluble fluoride can enter the root system by passive or active absorption, depending on its concentration. Fluoride at lower concentrations in solution (0.1–10 mg/L) is mainly absorbed and enriched in the root system of tea plants via active absorption, with a kinetic curve following the Michalis–Menten kinetic model. At higher concentrations (50–100 mg/L), water-soluble fluoride is absorbed at a rate that increases with increasing concentration, and this is achieved via passive absorption[17]. Many studies have shown that the water-soluble fluoride content in soils in most tea-producing regions in China is below the threshold for passive absorption[1820], indicating that fluoride mainly enters tea plants via active absorption by the roots.

    The active uptake of fluoride by tea roots is mediated by ion pump carrier proteins and ABC transporter proteins. Ion pump carrier proteins can transport the substrate across the cell membrane against the electrochemical gradient, and use the energy of ATP hydrolysis to participate in the process of active transport of substances, mainly including proton pump H+-ATPase and calcium ion pump Ca2+-ATPase. ABC transporter proteins can transport ions or heavy metals to vesicles as chelated peptide complexes, thereby reducing toxicity to the cell and improving plant resistance to abiotic stress. Passive fluoride absorption by the root system of tea plants involves water channels and ion channels. Studies on the effects of applying external anion channel inhibitors, cation channel inhibitors, and water channel inhibitors showed that inhibition of external anion channels significantly reduced the absorption of fluoride by roots. This indicated that anion channels are an important pathway for the uptake and trans-membrane transport of fluoride in the root system of tea plants[17,21,22]. The homeostatic flow of ions through channels diffusing along a trans-membrane concentration gradient or potential gradient involves ion channel proteins[23]. Two phylogenetically independent ion channel proteins have recently been identified in tea plants: CLCF-type F/H+ reverse transporter proteins and the FEX (Fluoride export gene) family of small membrane proteins[24]. CLC proteins are involved in the transport of a variety of anions such as chlorine (Cl) and F into and out of the cell. The FEX proteins in tea plants are involved in fluoride absorption via thermodynamic passive electro-diffusion through transmembrane channels[25,26].

    After fluoride is absorbed, it is transported within the tea plant by several different pathways. These include transport after leaf absorption, transport after root absorption, and transfer inside tea plant cells. The fluoride absorbed by leaves can be transferred along the conduit to the leaf tips and edges, accumulating in the top and ipsilateral leaves, but not in the roots. Fluoride in the soil and water environment is taken up by the root system and transported to the xylem via intracellular and intercellular transport. It is then transported upwards via transpiration and eventually accumulates in the leaves[27,28]. Two transport mechanisms have been proposed for the translocation of fluoride via the xylem. One proposed mechanism is that it is translocated in the form of fluoride-aluminum complexes[29]. The other proposed mechanism is that aluminum and fluoride are transported separately and accumulate after reaching the leaves[30].

    The roots are the main organ responsible for fluoride absorption in tea plants. Therefore, most studies have focused on the transport process in roots, especially the transport of fluoride by fluoride-related transporters. It has been found that under acidic conditions, F preferentially forms complexes with Al3+ and these complexes are then absorbed by roots and transported upward in the same state[29,31]. Studies have shown that, compared with F, aluminum-fluoride complexes are more easily absorbed and transported to the new shoots by the root system. This may be related to the elimination of the separate toxic effects of F and Al3+ [29]. Fluoride can also be transported in tea plants by binding to aluminum-organic acid complexes, and then accumulate in the leaves[29]. Both tea plant fluoride transporter proteins, CsFEX1 and CsFEX2, are involved in fluoride transport, but their encoding genes can be differentially expressed among different varieties and depending on the concentration of fluoride. In one study, the expression level of CsFEX1 was consistent among different varieties, while the expression of CsFEX2 was induced under fluoride stress to increase fluoride efflux from tea plants, thereby reducing its accumulation in low-fluoride varieties[32]. In addition, the A–G subfamily of ABC transporters plays a carrier role in the transmembrane transport of F- and Cl- in tea plants[33]. It was found that the expression of the ABC transporter protein CsCL667 was up-regulated in response to fluoride treatment, and its ability to transport fluoride was enhanced, suggesting that CsCL667 functions in fluoride efflux[34]. Another study demonstrated that CsABCB9 localizes in chloroplasts and functions as a fluoride efflux transporter to reduce fluoride-induced damage in leaves and enhance chloroplast activity[35].

    Several factors affect the absorption and transport of fluoride in tea plants, including the absorbable fluoride concentration, soil pH, the presence of other ions, and the activity of ion channels[3638]. Fluoride in nature is present in the atmosphere and soil, and its concentration is the main factor affecting the fluoride content in tea plants. Under normal conditions, tea plants generally absorb fluoride from the soil through the roots, but when the hydrogen fluoride content in the atmosphere is high, tea plants can absorb it through the leaves. The fluoride content in tea plants growing in the same geographical area is similar, mainly because of the soil properties in that area. Tea plants grow in acidic environments, and fluoride in acidic soils is more easily absorbed. During the growth of tea plants, the roots secrete organic acids such as oxalic acid, citric acid, and malic acid, which promote the absorption of fluoride by the roots and its transport to above-ground parts[39]. Other ions such as Ca2+ and Mg2+ combine with F to form precipitates, resulting in lower concentrations of water-soluble fluoride in the soil, which also affects its absorption by tea plant roots[40]. Exogenously applied calcium at low concentrations can change the cell wall structure and membrane permeability in tea plant roots, ultimately leading to reduced fluoride content in tea leaves[36,41,42]. Meanwhile, Al3+ treatment can trigger Ca2+ signaling in tea plant roots, which in turn activates calmodulin and promotes fluoride absorption[43]. The H+ gradient generated by the plasma membrane H+-ATPase can also promote Ca2+ signaling in plants to regulate the transmembrane transport of ions, which affects fluoride absorption[43]. The abundance and activity of H+-ATPase in the plasma membrane of tea plant roots have been found to increase significantly under fluoride stress, and these increases result in improved absorption of fluoride, although this is also affected by the fluoride concentration and temperature[33]. Sodium fluoride was found to induce the expression of genes encoding ABC transporter proteins, resulting in the transmembrane absorption of large amounts of fluoride ions into cells[34]. ABC transporters also transport ions alone or in the form of chelated peptide complexes directly out of the cellular membrane, which improves cellular tolerance to these ions[44]. Some anions with the same valence state also affect the F content in tea plants. For example, the ion channel protein encoded by CLCF is more sensitive to F, more selective for F than for Cl, and functions to export F from the cytoplasm to protect against fluorosis[4547].

    It can be seen that reducing the absorption of fluorine by tea plants and changing the mechanism of fluorine transportation in tea plants can reduce the content of fluorine in different parts of the tea plants. From the mechanism of fluorine absorption, the most effective way is to directly change the form of soil fluorine to reduce the absorption of water-soluble fluorine by roots. On this basis, it is possible to further change the active absorption process of fluorine mediated by ion pump carrier protein and ABC transporter protein in tea roots by molecular techniques. From the perspective of fluorine transport mechanism, the toxic effect of fluorine on tea plants can be reduced mainly by promoting the function of transporter proteins to exclude fluorine from the cell or transport it to the vesicle. The comprehensive application of the above methods to limit fluorine absorption and promote fluorine transport in tea plants can limit the accumulation of fluorine in tea plants.

    The fluoride enrichment characteristics of tea plants are related to various factors, including the tea variety, the organ, and the season. Among different varieties of tea, differences in leaf structure and other physiological characteristics can lead to variations in fluoride absorption and enrichment[48]. Some studies have concluded that the variety is one of the main determinants of the fluoride content in tea leaves[10], and the differences in fluoride content among most varieties reached highly significant levels (Table 1), which can be divided into low-enriched, medium-enriched, and high-enriched germplasm[49]. Various organs of tea plants also show differences in fluoride accumulation. The fluoride content is much higher in leaves than in roots and stems, and significantly higher in old leaves than in new shoots[10,50,51]. The fluoride content can differ widely among tea plants at different developmental stages. In spring, the new leaves begin to accumulate fluoride from the environment, and the fluoride content increases as the leaves age. When the growth rate of tea leaves is slower, they absorb and accumulate more fluoride from the soil and air. When the temperature in summer and autumn is high, the growth rate of tea leaves is fast and the growth period is short, so less fluoride is absorbed and accumulated from the soil and air. This explains why the fluoride content in fresh tea leaves was higher in spring and relatively lower in summer and autumn[4]. Another study found that, in China, the fluoride content in tea leaves was higher in summer than in spring. This may have been related to the maturity level of the tea leaves at harvest and different patterns of fluoride transport[4].

    Table 1.  Fluoride content difference of different tea cultivars.
    Cultivars Province Parts Treatment Years Content (mg/kg) Ref.
    Liannandaye Sichuan Old leaves Drying at 80 °C and boiling water extraction 2006−2007 1,150.79 ± 4.86 [107]
    Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 1,296.66 ± 12.84 [110]
    Yuenandaye Old leaves Drying at 80 °C and boiling water extraction 2006−2007 1,352.89 ± 12.69 [107]
    Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 1,560.36 ± 27.10 [110]
    Chenxi NO.4 Old leaves Drying at 80 °C and boiling water extraction 2006−2007 1,865.61 ± 7.46 [107]
    Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 1,954.93 ± 10.96 [110]
    Meizhan Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,180.13 ± 14.42 [107]
    Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 1,732.2 ± 41.2 [110]
    Zhejiang Mature leaves Drying at 80 °C and nitric acid extraction 2002 2,015.48 ± 29.99 [106]
    Fudingdabai Sichuan Old leaves Drying at 80 °C and boiling water extraction 2006−2007 249.64 ± 24.3 [107]
    Guizhou One bud and
    five leaves
    Drying at 80 °C and hydrochloric acid extraction 2010 1,612.3 ± 43.1 [109]
    Fudingdabai Zhejiang Mature leaves Drying at 80 °C and nitric acid extraction 2002 137.1 ± 2.1 [106]
    Fujian Old leaves Drying at 80 °C and hydrochloric acid extraction 2010 282.1 [111]
    Hunan One bud and
    five leaves
    Steaming and boiling water extraction 2011 2,232.05 ± 85.52 [51]
    Zhuyeqi Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,750.16 ± 11.37 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 125.4 [110]
    Hunan One bud and
    five leaves
    Steaming and boiling water extraction 2011 2,330.74 ± 31.39 [51]
    Fujianshuixian Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,548.18 ± 40.97 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 103.7 ± 1.5 [110]
    Fujian Old leaves Drying at 80 °C and hydrochloric acid extraction 2010 1,150.79 ± 4.86 [111]
    Huangyeshuixian Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,424.70 ± 18.85 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 2,950.80 ± 27.73 [110]
    Qianmei 701 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,522.01 ± 45.33 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,693.09 ± 35.12 [110]
    Guizhou One bud and
    five leaves
    Drying at 80 °C and hydrochloric acid extraction 2010 389.95 ± 32.18 [109]
    Guizhou Old leaves Dry samples and hydrochloric acid extraction 2011 2,142.26 ± 16.30 [113]
    Mingshan 130 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,564.78 ± 51.22 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,036.13 ± 31.25 [110]
    Mengshan 9 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,647.31 ± 70.89 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,436.55 ± 20.21 [110]
    Yinghong NO. 2 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,669.02 ± 799.95 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,364.53 ± 51.72 [110]
    Mengshan 11 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,695.21 ± 59.89 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,582.83 ± 9.73 [110]
    Mingshan 311 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,716.22 ± 42.21 [107]
    Donghuzao 2,731.20 ± 20.78
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,107.27 ± 54.91 [110]
    Hainandaye Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,746.82 ± 39.71 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 2,961.53 ± 29.94 [110]
    Qianmei 502 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,878.23 ± 76.94 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,881.51 ± 16.48 [110]
    Guizhou One bud and
    five leaves
    Drying at 80 °C and hydrochloric acid extraction 2010 389.95 ± 30.2 [109]
    Old leaves Dry samples and hydrochloric acid extraction 2011 3,260.48 ± 32.12 [113]
    Zisun Ya'an and
    surroundings
    Drying at 80 °C and boiling water extraction 2006−2007 2,904.13 ± 35.40 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,140.80 ± 42.86 [110]
    Zhejiang Drying at 80 °C and nitric acid extraction 2002 1,742.7 ± 43.2 [106]
    Qianmei 303 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,918.13 ± 46.79 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 4,029.11 ± 81.86 [110]
    Guizhou One bud and
    five leaves
    Drying at 80 °C and hydrochloric acid extraction 2010 199.74 ± 16.6 [109]
    Old leaves Dry samples and hydrochloric acid extraction 2011 2,972.79 ± 169.82 [113]
    Anxishuixian Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 2,924.33 ± 41.39 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,454.68 ± 26.29 [110]
    Longjing 43 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 3,152.73 ± 27.70 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 4,437.79 ± 26.14 [110]
    Zhejiang Drying at 80 °C and nitric acid extraction 2002 1,377.1 ± 37.0 [106]
    Fujian Old leaves Drying at 80 °C and hydrochloric acid extraction 2010 116.2 ± 0.9 [111]
    Shuyong 307 Ya'an and
    surroundings
    Drying at 80 °C and boiling water extraction 2006−2007 3,223.55 ± 151.43 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 4,296.52 ± 54.98 [110]
    Zhenghedabaicha Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 3,295.74 ± 27.55 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 3,876.58 ± 21.09 [110]
    Zhejiang Drying at 80 °C and nitric acid extraction 2002 1,373.0 ± 41.9 [106]
    Taiwandaye Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 3,363.59 ± 456.39 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 4,739.89 ± 58.59 [110]
    Qianmei 419 Ya'an and
    surroundings
    Old leaves Drying at 80 °C and boiling water extraction 2006−2007 3,518.15 ± 76.19 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 4,541.43 ± 28.91 [110]
    Guizhou One bud and
    five leaves
    Drying at 80 °C and hydrochloric acid extraction 2010 133.70 ± 11.2 [109]
    Old leaves Dry samples and hydrochloric acid extraction 2011 2,370.47 ± 11.43 [113]
    Mengshan23 Ya'an and
    surroundings
    Drying at 80 °C and boiling water extraction 2006−2007 3,625.11 ± 86.07 [107]
    Ya'an Mature leaves Drying at 70 °C and hydrochloric acid extraction 2006 4,469.25 ± 40.85 [110]
    Sichuan group species 2,782.59 ± 146.46
    Meitantaicha Guizhou One bud and
    five leaves
    Drying at 80 °C and hydrochloric acid extraction 2010 272.93 ± 27.3 [109]
    Qianmei 101 Drying at 80 °C and hydrochloric acid extraction 2010 175.07 ± 13.5
    Dry samples and hydrochloric acid extraction 2011 3,218.33 ± 57.91 [113]
    Qianmei 601 Drying at 80 °C and hydrochloric acid extraction 2010 267.11 ± 26.31 [109]
    Dry samples and hydrochloric acid extraction 2011 2,823.02 ± 73.36 [113]
    Qianmei 809 Drying at 80 °C and hydrochloric acid extraction 2010 521.48 ± 50.32 [109]
    Dry samples and hydrochloric acid extraction 2011 2,327.91 ± 83.17 [113]
    Qianmei 308 Drying at 80 °C and hydrochloric acid extraction 2010 326.88 ± 29.3 [109]
    Dry samples and hydrochloric acid extraction 2011 3,432.86 ± 159.4 [113]
    Qianmei 415 Drying at 80 °C and hydrochloric acid extraction 2010 186.95 ± 14.2 [109]
    Dry samples and hydrochloric acid extraction 2011 5,090.83 ± 69.56 [113]
    Qiancha NO. 7 Drying at 80 °C and hydrochloric acid extraction 2010 218.81 ± 18.7 [109]
    Qianfu NO. 4 136.82 ± 11.6
    Dry samples and hydrochloric acid extraction 2011 3,066.49 ± 86.35 [113]
    Guiyucha NO. 8 Drying at 80 °C and hydrochloric acid extraction 2010 191.03 ± 18.6 [109]
    Dry samples and hydrochloric acid extraction 2011 2,882.94 ± 195.73 [113]
    Pingyangtezao Drying at 80 °C and hydrochloric acid extraction 2010 125.02 ± 12.1 [109]
    Yuanxiaolv 244.32 ± 20.5
    Nongkangzao 133.70 ± 12.3
    Mingshan 213 106.98 ± 6.74
    Mingke NO. 4 195.29 ± 16.8
    Maolv 174.73 ± 15.8
    Qianmei 412 Old leaves Dry samples and hydrochloric acid extraction 2011 3,396.92 ± 31.61 [113]
    Meitantaicha 2011 3,085.83 ± 101.9
    Zhenong 138 Zhejiang Mature leaves Drying at 80 °C and nitric acid extraction 2002 805.7 ± 6.0 [106]
    Zhenong 12 1,041.2 ± 23.3
    Shuigu 1,123.2 ± 33.5
    Hanlv 1,152.4 ± 2.4
    Zhuzhichun 1,248.2 ± 2.3
    Lvyafoshou 1,298.1 ± 2.0
    Zhenong 139 1,322.4 ± 40.7
    Shuixian 1,323.5 ± 36.1
    Biyun 1,400.9 ± 0.6
    Soubei 1,487.6 ± 29.7
    Maoxie 1,487.7 ± 31.0
    Anhui NO. 9 1,489.7 ± 40.0
    Zhenong 113 1,492.7 ± 43.5
    Yingshuang 1,509.9 ± 7.2
    Zhenong 25 1,521.2 ± 3.2
    Ribenzhong 1,543.8 ± 33.9
    Yunqi 1,549.3 ± 46.9
    Zhenong 23 1,576.7 ± 11.3
    Huangyezao 1,606.7 ± 40.5
    Jinshi 1,662.4 ± 42.4
    Pingyun 1,676.6 ± 44.6
    Zhenong 21 1,678.8 ± 49.6
    Jinfeng 1,705.2 ± 10.3
    Jiukeng 1,779.2 ± 5.0
    Juhuachun 1,993.4 ± 14.5
    Wuniuzao 2,163.2 ± 15.8
    Fujian Old leaves Drying at 80 °C and hydrochloric acid extraction 2010 98.0 ± 1.3 [111]
    Jinguanyin All leaves Drying at 80 °C and nitric acid extraction 2014−2015 536.49 ± 10.41 [112]
    Dangui 2,598.87 ± 24.12
    Old leaves Drying at 80 °C and hydrochloric acid extraction 2010 145.3 ± 0.2 [111]
    Jinmudan All leaves Drying at 80 °C and nitric acid extraction 2014−2015 1,030.21 ± 36.52 [112]
    Ruixiang Drying at 80 °C and nitric acid extraction 1,315.64 ± 21.56
    Xiapu yuanxiao Old leaves Drying at 80 °C and hydrochloric acid extraction 2010 124.6 ± 3.0 [111]
    Jinxuan 103.7 ± 3.5
    Fuandabai 103.0 ± 1.0
    Fuyun NO. 7 104.0 ± 1.1
    Fuyun NO. 6 131.6 ± 1.8
    Fudingdahao 118.0 ± 2.4
    Zaochunhao 107.7 ± 2.8
    Xiapu chunbolv 99.2 ± 1.2
    BaijiguanF1 102.1 ± 1.1
    Huanguanyin Old leaves Drying at 80 °C and hydrochloric acid extraction 99.2 ± 1.3
    Foxiang NO. 1 Yunnan One bud and
    four leaves
    Drying at 60 °C and hydrochloric acid extraction 2011 155.00 ± 6.94 [108]
    Foxiang NO. 2 214.30 ± 5. 94
    Foxiang NO. 4 219. 50 ± 7. 32
    Foxiang NO. 5 190.70 ± 4.09
    Yunkang NO. 10 121. 30 ± 5. 81
    Yunkang NO. 14 198.50 ± 8.49
    Yuncha NO. 1 135.10 ± 4.74
    Baihaozao Hunan One bud and
    five leaves
    Steaming and boiling water extraction 2011 113.2 [51]
    Bixiangzao 121.4
    Taoyuandaye 177.7
    Yulv 165.9
    Jianbohuang NO.13 168.5
    Gaoyaqi 162.8
     | Show Table
    DownLoad: CSV

    In conclusion, selecting low-fluoride tea varieties and reducing the maturity of dark tea raw materials can be used as effective measures to reduce the fluoride content of dark tea.

    Plants can increase their resistance to fluoride through exocytosis and internal tolerance mechanisms. A series of reactions occurs in tea plants to reduce the toxic effects of fluoride and improve tolerance. The results of recent studies indicate that both external and internal factors are involved in fluoride resistance in tea plants. The external factors include the availability of cations that readily chelate fluoride, and the internal factors include the abundance and activity of certain transporters and the capacity of transporter and antioxidant systems (Fig. 3).

    Figure 3.  Mechanisms of fluoride tolerance in tea plants. (A) Metabolites in tea plants reduce the toxic effect of fluoride. Cell wall macromolecular components such as pectin, lignin, cellulose, hemicellulose, polysaccharide, and proteins chelate fluoride. The contents of some metabolites increase during adaptation of tea plants to fluoride. (B) Complexation of cations (Al3+, Ca2+, and Mg2+) with fluoride in tea plants. (C) Roles of the antioxidant system of tea plant in fluoride tolerance. Increases in the activity/abundance of antioxidant enzymes and in the ASA-GSH cycle reduce intracellular levels of reactive oxygen species, leading to increased fluoride tolerance. (D) Roles of transporters in fluoride tolerance. Transporter proteins transport fluoride into the vacuole, and this compartmentalization reduces damage to enzymes and organelles. The CsFEX, CsCLC, and CsABC transporters efflux fluoride from cells, thereby reducing its toxic effects. POD: Peroxidase, CAT: Catalase, SOD: Superoxide dismutase, APX: Ascorbate peroxidase, GR: Glutathione reductase, DHAR: Dehydroascorbate reductase, ABA: Abscisic acid, GA: Gibberellic acid, GHs: Glycoside hydrolases, ASA-GSH: Antioxidant system and the ascorbate-glutathione, ROS: Reactive oxygen species.

    Fluoride ions have a strong ability to form complexes with metal ions. Free F can form complexes with cations such as Al3+, Fe3+, and Ca2+, thereby altering ionic homeostasis and reducing its toxicity to tea plants[52]. Fluoride and aluminum ions form complexes and are enriched in leaves and other organs with a certain proportion. This reduces the toxicity of both Fl- and Al3+, and may be an important physiological mechanism of fluoride enrichment in tea plants[31,53]. Fluoride can also form complexes with Ca2+, so exogenous application of Ca2+ can effectively reduce the fluoride content and enhance the fluoride resistance of tea plants[42]. Fluoride combines with Mg2+, Al3+, and Ca2+ on the surface of tea leaves, and is present on the abaxial and adaxial leaf surfaces in the form of MgF2 and AlF3. The application of a small amount of MgF2 or CaF2 may be a means to reduce the toxicity of fluoride to tea seedlings[40]. Treatment with selenium was shown to reduce the fluoride content in tea, increase the accumulation of fluoride in roots, and reduce the proportion of water-soluble fluoride in tea beverages[54].

    The cell wall is widely involved in plant growth and development and in various stress responses. Fluoride ions can be chelated by the aldehyde, carboxyl, amino, and phosphate groups in polysaccharides, pectin, lignin, proteins, and other components as well as some metal ions adsorbed in the cell wall, which is usually called cell wall fixation[55,56]. Recent studies have shown that fluoride stress activates pathways related to cell wall metabolism, the stress response, signal transduction, and protein degradation, and all of these pathways may contribute to the accumulation/detoxification of fluoride in tea leaves[57,58]. After the application of exogenous fluoride, there are increase in the activities of key enzymes involved in the pectin biosynthetic pathway, in the transcript levels of their encoding genes, and in the pectin polysaccharide content, indicating that treatment with exogenous fluoride promotes pectin biosynthesis. In turn, it promotes the combination of absorbed fluoride with pectin[59]. Lignin is the main component of the plant cell wall, and its amount and the activity of its biosynthetic pathway increase in response to fluoride stress. The lignin content showed the same trend as the fluoride content in leaves, consistent with its important role in alleviating fluoride toxicity in tea plants[15]. Tea polysaccharides can also adsorb and bind fluoride. Compared with polysaccharides in other plants, tea polysaccharides have the highest fluoride content and the strongest fluoride complexation ability. The majority (80%) of fluoride in tea is bound with tea polysaccharides, and the formation of these complexes is one of the factors that enhances tea plants’ fluoride resistance[60]. Studies have found that with increasing fluoride concentration, the F content in the cell wall and its components, the metal ion content in the cell wall, and the contents in total cell wall materials, cellulose, and pectin increased with highly significant positive correlations[59,61].

    To maintain normal plant growth under adverse conditions, a series of metabolic reactions occur to activate defense responses. The formation and transformation of secondary metabolites under fluoride stress may be one way in which tea plants resist fluoride. One study found that as the fluoride concentration increased, catechin was catabolized to produce lignin, the polyphenol content decreased, and the lignin content increased. Thus, leaf lignification promotes stress resistance in tea plants[62]. Organic acids, carbohydrates, and amino acids also play important roles in the fluoride tolerance of tea plants[63]. The contents of free proline and citric acid were found to increase under fluoride stress, and the oxalic acid content in leaves first increased and then decreased as the fluoride concentration increased. These patterns of accumulation suggested that these metabolites were involved in a protective response against fluoride stress in tea plants[64,65]. Another study detected up-regulation of glycoside hydrolases (GHs), primary amine oxidase, and citrate synthetase under fluoride stress, indicating that these enzymes may be involved in the defense response[66]. Plant growth regulators such as abscisic acid and gibberellin play important roles in the response to fluoride stress and in signal transduction[67,68]. However, further studies are required to explore the roles of these and other plant growth regulators in the responses to fluoride stress and in fluoride enrichment in tea plants.

    Subcellular distribution analyses in tea plants have shown that F is concentrated in vacuoles in the cells of tea leaves, indicating that vacuoles are the main site of fluoride accumulation. Fluoride transporters are involved in the vacuole sequestration of fluoride[52]. The fluoride transporter gene FEX in tea plants is expressed in a tissue-specific manner and its product can enhance tolerance to fluoride by reducing the fluoride content in tissues[26]. Studies have shown that fluoride treatment activates the expression of genes encoding receptor-like kinases and MYB and MADS-box transcription factors, thereby regulating fluoride accumulation and fluoride tolerance in tea plants[6971]. Exploring the regulatory mechanism of fluoride transporters is the key to understanding the fluoride enrichment characteristics of tea plants, and is a new direction for molecular research.

    Under fluoride stress, tea plants can eliminate excess reactive oxygen species (ROS) within a certain concentration threshold by regulating their metabolism, thereby protecting themselves against oxidative damage. Under low-level or short-term fluoride treatments, the antioxidant system and the ascorbate-glutathione (ASA-GSH) cycle respond to fluoride stress, and there are increase in the activities of glutathione reductase, ascorbate peroxidase, dehydroascorbate reductase, peroxidase, catalase, and superoxide dismutase. Together, these enzymes remove ROS to reduce the toxicity of fluoride to tea plants. Tea plants that accumulate high levels of fluoride show a stronger ability to remove ROS[72]. Selenium treatment can also modulate fluoride-induced oxidative damage by increasing the activities of superoxide dismutase, peroxidase, and catalase, resulting in reduced malondialdehyde levels[54]. However, as the fluoride concentration increases beyond the detoxification capacity of protective enzymes and non-enzymatic antioxidants in both systems, ROS accumulate to excess levels and cause damage to tea plants[73].

    In conclusion, there are different forms of resistance to fluoride stress in the tea plants, so the adaptability of tea plants to fluorine stress can be improved by enhancing these resistance mechanisms. Appropriate agronomic measures in the tea gardens can enhance the expression level of stress resistance genes in the tea plants, and then increase the content of downstream metabolites to enhance stress resistance. At the same time, the necessary molecular technology can be used as an auxiliary means to carry out a certain aspect of the targeted improvement, and comprehensively enhance the fluorine tolerance of tea plants.

    Fluoride has dual effects on tea plant growth and metabolites related to tea quality. At low concentrations, fluoride has no obvious effect on growth and can promote the normal physiological metabolism of tea plants. High concentrations of fluoride adversely affect tea plants, inhibit growth, and exert toxic effects to reduce the yield. In addition to yield, metabolites related to tea quality are affected by fluoride. At high concentrations, fluoride reduces the synthesis of key secondary metabolites, free amino acids, polyphenols, and caffeine in tea plants, thereby reducing tea quality. In addition, leaf materials with a high fluoride content result in tea beverages with a high fluoride content (Fig. 4). In this way, excessive fluoride seriously affects the quality and safety of tea. Long-term drinking of tea with a high fluoride content can cause skeletal fluorosis, which endangers the health of consumers.

    Figure 4.  Effects of fluoride stress on tea plant growth and tea quality. Fluoride at low concentrations increases the chlorophyll content, photosynthetic rate, and quality-related metabolites in tea plants; and increases the activity of the antioxidant system and the ASA-GSH cycle to remove reactive oxygen species (ROS). Fluoride at high concentrations that exceed the tolerance limit of tea plants decreases the scavenging capacity of the antioxidant system and the ASA-GSH cycle, resulting in ROS accumulation. In addition, damage to chloroplast thylakoid membranes and decreases in chlorophyll content lead to decreases in the photosynthetic rate, stomatal conductance, and carbon assimilation capacity, resulting in decreased biomass and decreased content of quality-related metabolites, as well as leaf yellowing, leaf abscission, and even plant death. POD: Peroxidase, CAT: Catalase, SOD: Superoxide dismutase, APX: Ascorbate peroxidase, GR: Glutathione reductase, DHAR: Dehydroascorbate reductase, ASA-GSH: Antioxidant system and the ascorbate-glutathione, ROS: Reactive oxygen species.

    The growth responses of tea plants to fluoride depend on its concentration. When tea plants were treated with a low concentration of fluoride, the chlorophyll content and photosynthetic rate increased slightly, the initial respiration mode shifted from the glycolysis pathway to the pentose phosphate pathway, and respiration was enhanced. When tea plants were treated with a high concentration of fluoride, the toxic effect was mainly manifested as inhibition of metabolism and damage to cell structure. Excessive fluoride can damage the chloroplasts and cell membrane system of plants. Fluoride can also combine with Mg2+ in chlorophyll, resulting in damage to the chloroplast thylakoid membranes and significant decreases in the leaf photosynthetic rate, chlorophyll content, net photosynthetic rate, and stomatal conductance[7375]. Fluoride can also inhibit the carbon assimilation process by inhibiting the activity of rubisco, and inhibit the activity of ATP synthase on the thylakoid membrane of chloroplasts, thus hindering photophosphorylation[76]. Fluoride significantly inhibits the activities of enzymes involved in respiration, and causes the mitochondria of tea leaves to become vacuolated and degraded. In severe cases, it causes irreversible damage to mitochondria, which in turn leads to a smaller surface area for enzymes to attach to, resulting in weakened cellular respiration. Blocking of sugar metabolism in tea plants reduces respiration, and so ROS accumulate to excess levels[77,78]. Therefore, fluoride at high concentrations can lead to dwarfism, reduced growth, and leaf chlorosis[79,80]. However, few studies have explored the mechanism of fluoride’s effect on tea plant photosynthesis and respiration, and further research is needed.

    Metabolites that contribute to tea quality include polyphenols, amino acids, alkaloids, and aroma substances. The main class of polyphenols is catechins, followed by flavonoids and anthocyanins. Tea polyphenols confer astringency, an important taste quality character. In addition, the oxidation products of tea polyphenols such as theaflavins and thearubigins contribute to the infusion color of fermented teas such as black tea. Most of the flavonols of tea polyphenols are combined with a glycoside to form flavonoid glycosides, which are important contributors to the infusion color of non-fermented teas such as green tea[81]. Tea polyphenols are important antioxidants, and have tumor-inhibiting, anti-inflammatory, and antibacterial activities. Amino acids contribute to the freshness of tea infusions and are an important tea quality parameter. Amino acids can be divided into protein-source amino acids and non-protein-source amino acids. Theanine, a non-protein-source amino acid, is the main amino acid in tea. Theanine contributes to the freshness of tea infusions and offsets the astringency and bitterness of catechin and caffeine. It also has the effect of calming the nerves and promoting sleep in humans[82,83]. The main alkaloid in tea plants is caffeine, which is mainly synthesized and stored in the leaves, and is often stored in the vacuole as a complex with chlorogenic acid. Caffeine affects the quality of tea infusions, and contributes to the bitter taste. It also forms complexes with theaflavins and other substances with a refreshing taste. The quality of tea products is generally positively correlated with the caffeine content[84]. Caffeine has a stimulating effect and promotes blood circulation. Aroma substances in tea confer its unique scent and are important tea quality characters. The aroma of tea is not only an important and pleasant sensory character, but also an important factor in promoting human health.

    Previous studies have shown that fluoride treatments lead to changes in the types and abundance of metabolites such as minor polypeptides, carbohydrates, and amino acids in tea. However, depending on its concentration, fluoride can have dual effects on the physiological metabolism of tea plants. The contents of tea polyphenols, amino acids, caffeine, and water extracts were found to be enhanced by low concentrations of fluoride, but inhibited by fluoride at high concentrations[85]. Similarly, a low-concentration of fluoride was found to increase the contents of the main aroma components in tea and improve tea quality[86,87]. A high fluoride concentration can lead to significant decreases in the amounts of some tea polyphenols, total catechins, protein, theanine, and caffeine, resulting in decreased tea quality[8789]. Aroma is an important quality character of tea, and studies have shown that the amounts of aroma compounds in tea decrease as the fluoride concentration increases. Most aroma compounds show a trend of increasing and then decreasing as the fluoride concentration increases, and only alcohols show the opposite trend. Thus, a high concentration of fluoride adversely affects tea aroma and flavor quality[86,88]. In general, a high fluoride concentration decreases the abundance of important quality metabolites such as tea polyphenols, amino acids, caffeine, and aroma substances, resulting in weakened taste intensity, freshness, and aroma quality. The quality formation of tea is extremely unfavorable under high-fluoride conditions.

    In summary, fluorine stress affects the growth and metabolite content of tea plants, and then affect the safety and quality of tea products. It is necessary to find suitable measures to reduce fluorine in tea garden production, which can increase the content of tea quality metabolites while ensuring or promoting the growth and development of tea plants. On this basis, it is worth studying to further enhance the content of fluoride-tolerant metabolites of tea plants and is a worthwhile research direction.

    Although tea plants have characteristics of polyfluoride and fluoride resistance, excessive fluoride accumulation can still impair their growth and affect tea yield and quality. The long-term consumption of dark tea made from thick, mature leaves can cause tea-drinking fluorosis, and so dark tea has become an important target of tea safety risk research. On the whole, screening for low-fluoride tea varieties, improving soil management measures in tea plantations, and improving tea processing technologies will contribute to reducing the fluoride content in tea and ensuring its quality and safety (Fig. 5).

    Figure 5.  Defluoridation measures for tea plants. (A) Breeding low-fluoride varieties of tea plants. (B) Improving management measures during tea plant cultivation. (C) Improving tea processing technologies. (D) Appropriate brewing methods to prepare tea infusions.

    The fluoride accumulation characteristics vary among tea varieties and are mainly controlled by genotype. Different tea varieties have different fluoride accumulation capabilities. Selecting appropriate low-fluoride varieties is the primary measure to reduce the fluoride content in tea[10]. The differences in fluoride content among varieties are related to differences in leaf structure. Large and thin leaves with well-developed spongy tissue and large intercellular spaces are conducive to absorbing fluoride from the atmosphere, and accumulate a higher fluoride content[86]. Tea plants mainly absorb fluoride through their roots, and there is a significant correlation between root activity and fluoride content in tea plant roots. Therefore, differences in root activity among varieties may explain differences in fluoride uptake. Studies have also shown that fluoride accumulation in tea plants may be affected by the branching angle, a character that is under moderate to strong genetic control[65]. The fluoride content varies widely among different varieties of tea. Breeding and cultivating tea varieties with low fluoride content is an effective way to produce tea beverages with low fluoride concentrations.

    Tea plants can absorb fluoride from the environment. The origin of tea plants and environmental factors directly affect the accumulation of fluoride[90]. Areas where there is a low fluoride content in the soil should be selected for the cultivation of tea plants. The irrigation water should be low-fluoride water, and there should be no fluoride pollution in the air. At the same time, improving soil management measures can effectively reduce the fluoride content in tea leaves. The use of phosphorus fertilizers should be reduced during the planting process, and chemical or organic fertilizers with low fluoride contents should be used to prevent soil pollution. The application of nitrogen fertilizers at appropriate levels combined with root fertilization and foliar spraying can also affect fluoride enrichment in tea plants[91]. Calcium in different forms and concentrations can form CaF2 with fluoride or change the surface charge of soil particles, ion exchange capacity, and the stability of complexes. These changes can alter the soil pH and affect the soil exchangeable fluoride content[37,92]. Competitive adsorption and material chelation reactions in the soil can reduce the absorbable fluoride content. The addition of charcoal from bamboo and other materials can significantly reduce the water-soluble and available fluoride content in tea garden soil, as well as increasing the contents of organically bound fluoride and Fe/Mn-bound fluoride. This method can reduce the absorption and accumulation of fluoride in tea plants without adversely affecting the contents of major secondary metabolites[93,94]. Humic acid aluminum (HAA) adsorbents and low-molecular-weight organic acids can significantly reduce the fluoride content in the soil solution by chelating soluble fluoride, ultimately reducing its absorption by tea plants[95]. Soil defluoridation agents in tea gardens can decrease the soil fluoride content[96], although they do not necessarily decrease the fluoride content in fresh tea leaves.

    The fluoride content in tea mainly depends on the fluoride content in fresh tea leaves, which is affected by the tea genotype and the soil environment. The processing method has a smaller effect on the fluoride content in tea beverages. Compared to green tea, white tea, black tea, yellow tea and oolong tea, the processing of dark tea uses more mature leaves and old leaves, which affects the fluorine content of the finished tea, and there is a risk of excessive fluorine content, so it is necessary to improve dark tea processing technologies to reduce its fluoride content. One study found that appropriate blending of tea raw materials is an effective processing method to reduce the fluoride content in tea leaves. In this method, the fluoride content is measured when selecting raw materials, and fresh tea leaves with different fluoride contents are screened. Blending raw materials with high fluoride content, medium fluoride content, and low fluoride content can effectively control the final fluoride content[39]. During processing, the fluoride content in the tea leaves can be effectively reduced by washing the rolled tea leaves with room-temperature water for 1–2 min, a process that retains the effective components to the greatest extent[97]. Before the dark tea fermentation process, spraying microbial agents while stirring can effectively reduce the fluoride content and improve the quality, aroma, and taste of dark tea. In the processes of tea manufacturing and deep processing, adding different defluorination agents can effectively reduce the fluoride content in tea products without affecting the quality[94,98,99]. Studies have found that Eurotium cristatum is able to phagocytose fluoride. The fluoride content in black tea was effectively reduced using a E. cristatum strain mutagenized by ultraviolet radiation[100].

    Tea beverages are generally prepared by brewing or boiling. The leaching rate of fluoride from tea is affected by factors such as the extraction time, extraction method, and brewing time. Therefore, the brewing method can affect fluoride intake. The fluoride content in matcha depends, in part, on the brewing conditions[9]. The fluoride leaching rate of dark tea was found to be significantly correlated with the brewing method, and was significantly higher in tea prepared using the boiling method than in tea prepared using the ordinary brewing method. The rate of fluoride leaching from tea prepared using the boiling method was also higher with tap water than with pure water. A higher ratio of tea to water, increased water temperatures, and prolonged brewing time also increase the leaching rate of fluoride from tea[101-103]. Therefore, to significantly reduce the intake of fluoride by consumers and prevent fluorosis, it is recommended that tea should be prepared using pure water for brewing, an appropriate tea-water ratio and water temperature, and a shorter brewing time. Adding food-grade nutritional supplements to tea infusions can also reduce the fluoride content below the standard, and does not significantly affect the other bioactive components and quality factors[98].

    The issue of tea safety is an important concern in society. Research on the mechanisms of fluoride enrichment in tea plants and related research on fluoride control and defluorination technologies is of great significance to tea quality and safety, as well as tea genetics and breeding. Recently, some progress has been made in research on the fluoride enrichment and tolerance mechanisms of tea plants, and this has provided a theoretical basis for further research on methods to reduce the fluoride content in tea.

    (1) Although there has been some progress in research on how tea plants adsorb and transport fluoride, the specific mechanisms are still unclear. Further studies should focus on the molecular mechanisms of fluoride ion transport channel proteins (CLC, FEX, and ABC transporters), their interacting proteins, and how they are regulated to control fluoride enrichment. Studies have shown that the deposition of aluminum-fluoride complexes on the cell wall and compartmentalization in the vacuoles are important mechanisms for the detoxification of these ions in tea plants. However, it is still unclear which proteins regulate the absorption, efflux, transport, and storage of these complexes.

    (2) Tea is rich in secondary metabolites such as polyphenols, polysaccharides, and organic acid. Tea polysaccharides can combine with F to form complexes, thereby reducing the toxic effects of F ions[60]. Tea polyphenols contain multiple phenolic hydroxyl groups, have a strong acid-base buffering capacity, and can form complexes with various metal ions to generate ring-shaped chelates. The flavonol content of polyphenols was found to be significantly positively correlated with Al3+ accumulation, and their binding capacity was found to be higher than that of epigallocatechin gallate and proanthocyanidins in the root[104]. Whether polyphenols can further react with F after complexing with Al3+ is worthy of further study. It will be interesting to explore the roles and mechanisms of secondary metabolites in fluoride enrichment and tolerance in tea plants.

    (3) The degree of fluoride stress affects the growth and quality of tea[105]. How to maintain the balance between fluoride content and quality is a problem that needs to be solved in the industry. The reason why dark tea selects leaves with higher maturity is mainly because, in the same amount of leaves, the leaves with higher maturity contain more effective ingredients such as tea polyphenols, amino acids, trace elements and fiber required by the human body, and the finished dark tea with higher leaf maturity has a lower price and is more acceptable to consumers. If the fluorine content of dark tea is reduced by reducing the maturity of the raw materials, the taste will be inappropriate and difficult to be accepted by consumers. Therefore, it is necessary to reduce the fluorine content while maintaining the quality of dark tea, which needs further research .

    (4) There is still a lack of practical and effective fluoride reduction measures in the tea industry, and the development of such measures will be a key breakthrough. In terms of reducing fluoride levels in tea, the first step is to compare fluoride contents among different tea varieties and select varieties with relatively low fluoride content. The next steps are to improve the management of soil in tea plantations, improve tea processing technologies, and recommend appropriate brewing methods. However, there are still no systematic, efficient, and fully effective management measures for reducing the fluoride content in tea. Breeding new low-fluoride varieties of tea plants using traditional breeding methods is long and difficult, and has not yet been achieved using modern molecular breeding technologies. The use of a single defluoridation measure has certain limitations, so it is advisable to combine several strategies to reduce the fluoride content in tea leaves.

    The authors confirm contribution to the paper as follows: study conception and design: Zeng L; data collection: Yang J, Liu C; analysis and interpretation of results: Yang J, Liu C, Zeng L; draft manuscript preparation: Yang J, Liu C, Li J, Zhang Y, Zhu C, Gu D, Zeng L. All authors reviewed and approved the final version of the manuscript.

    The datasets generated during and/or analyzed during the current study are not publicly available due to management requests, but are available from the corresponding author on reasonable request.

    Part of the research aspects carried out by the authors are supported by the financial support from the Key-Area Research and Development Program of Guangdong Province (2023B0202120001), the Guangdong Natural Science Foundation for Distinguished Young Scholar (2023B1515020107), Tea garden standardized production and processing project of Yigong tea farm in Nyingchi City, the South China Botanical Garden, Chinese Academy of Sciences (QNXM-202302), the fund for China Agriculture Research System (CARS-19), Chinese Academy of Sciences Specific Research Assistant Funding Program (2021000064, 2023000030), the Science and Technology Project of Guangzhou (202206010185), the Guangdong Provincial Special Fund for Modern Agriculture Industry Technology Innovation Teams (2023KJ120), and the Science and Technology plan Project of Qingyuan (220804107510735).

  • The authors declare that they have no conflict of interest.

  • [1]

    Abbasi Khalaki M, Moameri M, Asgari Lajayer B, Astatkie T. 2021. Influence of nano-priming on seed germination and plant growth of forage and medicinal plants. Plant Growth Regulation 93:13−28

    doi: 10.1007/s10725-020-00670-9

    CrossRef   Google Scholar

    [2]

    Reed RC, Bradford KJ, Khanday I. 2022. Seed germination and vigor: ensuring crop sustainability in a changing climate. Heredity 128:450−59

    doi: 10.1038/s41437-022-00497-2

    CrossRef   Google Scholar

    [3]

    Feeley KJ, Bravo-Avila C, Fadrique B, Perez TM, Zuleta D. 2020. Climate-driven changes in the composition of New World plant communities. Nature Climate Change 10:965−70

    doi: 10.1038/s41558-020-0873-2

    CrossRef   Google Scholar

    [4]

    Mestanza-Ramón C, Henkanaththegedara SM, Vásconez Duchicela P, Vargas Tierras Y, Sánchez Capa M, et al. 2020. In-Situ and Ex-Situ Biodiversity conservation in ecuador: a review of policies, actions and challenges. Diversity 12:315

    doi: 10.3390/d12080315

    CrossRef   Google Scholar

    [5]

    Turner SR, Steadman KJ, Vlahos S, Koch JM, Dixon KW. 2013. Seed treatment optimizes benefits of seed bank storage for restoration-ready seeds: the feasibility of prestorage dormancy alleviation for mine-site revegetation. Restoration Ecology 21:186−92

    doi: 10.1111/j.1526-100X.2012.00879.x

    CrossRef   Google Scholar

    [6]

    Lee SY, Park K, Jang BK, Ji B, Lee H, et al. 2022. Exogenous gibberellin can effectively and rapidly break intermediate physiological dormancy of Amsonia elliptica seeds. Frontiers in Plant Science 13:1043897

    doi: 10.3389/fpls.2022.1043897

    CrossRef   Google Scholar

    [7]

    Kaur H, Nazir F, Hussain SJ, Kaur R, Rajurkar AB, et al. 2023. Gibberellic acid alleviates cadmium-induced seed germination inhibition through modulation of carbohydrate metabolism and antioxidant capacity in Mung bean seedlings. Sustainability 15:3790

    doi: 10.3390/su15043790

    CrossRef   Google Scholar

    [8]

    Sadeghi F, Sohrabi Y, Mardeh ASS. 2023. Effects of plant growth regulators on seed germination and biochemical properties of two wheat cultivars under water deficit conditions. Gesunde Pflanzen 75:1121−32

    doi: 10.1007/s10343-022-00803-2

    CrossRef   Google Scholar

    [9]

    Vishal B, Kumar PP. 2018. Regulation of seed germination and abiotic stresses by gibberellins and abscisic acid. Frontiers Plant Science 9:368905

    doi: 10.3389/fpls.2018.00838

    CrossRef   Google Scholar

    [10]

    Singh D, Mishra M, Yadav A. 2016. Standardizing the methods for breaking seed dormancy to enhance germination of Gloriosa Superba Seeds. Expert Opinion on Environmental Biology 5:1

    doi: 10.4172/2325-9655.1000123

    CrossRef   Google Scholar

    [11]

    Kucera B, Cohn MA, Leubner-Metzger G. 2005. Plant hormone interactions during seed dormancy release and germination. Seed Science Research 15:281−307

    doi: 10.1079/SSR2005218

    CrossRef   Google Scholar

    [12]

    Dunlap JR, Morgan PW. 1977. Reversal of Induced Dormancy in Lettuce by Ethylene, Kinetin, and Gibberellic Acid. Plant Physiology 60:222−24

    doi: 10.1104/pp.60.2.222

    CrossRef   Google Scholar

    [13]

    Unterholzner SJ, Rozhon W, Papacek M, Ciomas J, Lange T, et al. 2015. Brassinosteroids are master regulators of gibberellin biosynthesis in Arabidopsis. The Plant Cell 27:2261−72

    doi: 10.1105/tpc.15.00433

    CrossRef   Google Scholar

    [14]

    Hong YF, Ho THD, Wu CF, Ho SL, Yeh RH, et al. 2012. Convergent starvation signals and hormone crosstalk in regulating nutrient mobilization upon germination in cereals. The Plant Cell 24:2857−73

    doi: 10.1105/tpc.112.097741

    CrossRef   Google Scholar

    [15]

    Dalal NV, Rai VR. 2004. In vitro propagation of Oroxylum indicum Vent. a medicinally important forest tree. Journal of Forest Research 9:61−65

    doi: 10.1007/s10310-003-0055-x

    CrossRef   Google Scholar

    [16]

    Mangena P. 2021. Analysis of correlation between seed vigour, germination and multiple shoot induction in soybean (Glycine max L. Merr.). Heliyon 7:e07913

    doi: 10.1016/j.heliyon.2021.e07913

    CrossRef   Google Scholar

    [17]

    Baskin JM, Baskin CC. 2004. A classification system for seed dormancy. Seed Science Research 14:1−16

    doi: 10.1079/SSR2003150

    CrossRef   Google Scholar

    [18]

    Yildiz M, Beyaz R, Gursoy M, Aycan M, Koc Y, et al. 2017. Seed Dormancy. In Advances in Seed Biology, eds. Jimenez-Lopez JC. Rijeka, UK: IntechOpen. pp. 85-101. doi: 10.5772/intechopen.70571

    [19]

    Debeaujon I, Léon-Kloosterziel KM, Koornneef M. 2000. Influence of the testa on seed dormancy, germination, and longevity in Arabidopsis. Plant Physiology 122:403−414

    doi: 10.1104/pp.122.2.403

    CrossRef   Google Scholar

    [20]

    Baskin JM, Baskin CC. 2021. The great diversity in kinds of seed dormancy: a revision of the Nikolaeva–Baskin classification system for primary seed dormancy. Seed Science Research 31:249−277

    doi: 10.1017/S096025852100026X

    CrossRef   Google Scholar

    [21]

    Le Roux LG, Robbertse PJ. 1997. Aspects relating to seed production in Gloriosa superba L. South African Journal of Botany 63:191−97

    doi: 10.1016/S0254-6299(15)30743-2

    CrossRef   Google Scholar

    [22]

    Mahajan YA, Shinde BA, Torris A, Gade AB, Patil VS, et al. 2023. Pre-Sowing Treatments, Seed Components and Water Imbibition Aids Seed Germination of Gloriosa superba. Seeds 2:15−29

    doi: 10.3390/seeds2010002

    CrossRef   Google Scholar

    [23]

    Rodrigues CR, Rodrigues BF. 2014. Enhancement of seed germination in Macaranga peltata for use in tropical forest restoration. Journal of Forestry Research 25:897−901

    doi: 10.1007/s11676-014-0536-0

    CrossRef   Google Scholar

    [24]

    Poobathy R, Zakaria R, Murugaiyah V, Subramaniam S. 2019. Surface sterilization and micropropagation of Ludisia discolor. Biocatalysis and Agricultural Biotechnology 22:101380

    doi: 10.1016/j.bcab.2019.101380

    CrossRef   Google Scholar

    [25]

    Hesami M, Daneshvar MH, Yoosefzadeh-Najafabadi M. 2018. Establishment of a protocol for in vitro seed germination and callus formation of Ficus religiosa L., an important medicinal plant. Jundishapur Journal of Natural Pharmaceutical Products 13:e62682

    doi: 10.5812/jjnpp.62682

    CrossRef   Google Scholar

    [26]

    Davoudpour Y, Schmidt M, Calabrese F, Richnow HH, Musat N. 2020. High-resolution microscopy to evaluate the efficiency of surface sterilization of Zea mays seeds. PLOS ONE 15:e0242247

    doi: 10.1371/journal.pone.0242247

    CrossRef   Google Scholar

    [27]

    Teixeira da Silva JA, Winarto B, Dobránszki J, Cardoso JC, Zeng S. 2016. Tissue disinfection for preparation of Dendrobium in vitro culture. Folia Horticulturae 28:57−75

    doi: 10.1515/fhort-2016-0008

    CrossRef   Google Scholar

    [28]

    Hesami M, Naderi R, Yoosefzadeh-Najafabadi M. 2018. Optimizing sterilization conditions and growth regulator effects on in vitro shoot regeneration through direct organogenesis in Chenopodium quinoa. BioTechnologia 99:49−57

    doi: 10.5114/bta.2018.73561

    CrossRef   Google Scholar

    [29]

    Mosoh DA, Khandel AK, Verma SK, Vendrame WA. 2023. Effects of sterilization methods and plant growth regulators on in vitro regeneration and tuberization in Gloriosa superba (L.). In Vitro Cellular & Developmental Biology-Plant 59:792−807

    doi: 10.1007/s11627-023-10387-9

    CrossRef   Google Scholar

    [30]

    Vendrame WA, Xu J, Beleski DG. 2023. Micropropagation of Brassavola nodosa (L.) Lindl. using SETISTM bioreactor. Plant Cell, Tissue and Organ Culture 153:67−76

    doi: 10.1007/s11240-022-02441-y

    CrossRef   Google Scholar

    [31]

    Raina R, Gupta LM. 1997. Increasing seed yield in glory lily (Gloriosa superba) - experimental approaches. Acta Horticulturae 502:175−80

    doi: 10.17660/ActaHortic.1999.502.27

    CrossRef   Google Scholar

    [32]

    Patel AI, Desai BS, Chaudhari BN, Vashi JM. 2020. Genetic improvement in Glory lily (Gloriosa superba L.): a review. International Journal of Chemical Studies 8:255−60

    doi: 10.22271/chemi.2020.v8.i4d.9701

    CrossRef   Google Scholar

    [33]

    Pickens KA, Affolter JM, Wetzstein HY, Wolf JHD. 2003. Enhanced seed germination and seedling growth of Tillandsia eizii in vitro. HortScience 38:101−4

    doi: 10.21273/HORTSCI.38.1.101

    CrossRef   Google Scholar

    [34]

    Mosoh DA, Prakash O, Khandel AK, Vendrame WA. 2024. Preserving Earth’s flora in the 21st Century: Climate, Biodiversity, and Global Change Factors (GCFs) since the mid-1940s. Frontiers in Conservation Science 5:1383370

    doi: 10.3389/fcosc.2024.1383370

    CrossRef   Google Scholar

    [35]

    Anandhi S, Rajamani K. 2012. Studies on seed germination and growth in Gloriosa superba L. Global Journal of Research on Medicinal Plants & Indigenous Medicine 1:524

    Google Scholar

    [36]

    Sen MK, Jamal M, Nasrin S. 2013. Sterilization factors affect seed germination and proliferation of Achyranthes aspera cultured in vitro. Environmental and Experimental Biology 11:119−23

    Google Scholar

    [37]

    Barampuram S, Allen G, Krasnyanski S. 2014. Effect of various sterilization procedures on the in vitro germination of cotton seeds. Plant Cell, Tissue and Organ Culture 118:179−85

    doi: 10.1007/s11240-014-0472-x

    CrossRef   Google Scholar

    [38]

    Gu M, Li Y, Jiang H, Zhang S, Que Q, et al. 2022. Efficient in vitro sterilization and propagation from stem segment explants of Cnidoscolus aconitifolius (Mill.) I. M. Johnst, a multipurpose woody plant. Plants 11:1937

    doi: 10.3390/plants11151937

    CrossRef   Google Scholar

    [39]

    Hashim SN, Ghazali SZ, Sidik NJ, Chia-Chay T, Saleh A. 2021. Surface sterilization method for reducing contamination of Clinacanthus nutans nodal explants intended for in-vitro culture. E3S Web of Conferences 306:01004

    doi: 10.1051/e3sconf/202130601004

    CrossRef   Google Scholar

    [40]

    Miché L, Balandreau J. 2001. Effects of rice seed surface sterilization with hypochlorite on inoculated Burkholderia vietnamiensis. Applied and Environmental Microbiology 67:3046−52

    doi: 10.1128/AEM.67.7.3046-3052.2001

    CrossRef   Google Scholar

    [41]

    Sauer DB, Burroughs R. 1986. Disinfection of seed surfaces with sodium hypochlorite. Phytopathology 76:745−49

    doi: 10.1094/Phyto-76-745

    CrossRef   Google Scholar

    [42]

    Hesami M, Daneshvar MH, Lotfi-Jalalabadi A. 2017. Effect of sodium hypochlorite on control of in vitro contamination and seed germination of Ficus religiosa. Iranian Journal of Plant Physiology 7:2157−62

    Google Scholar

    [43]

    Ahsan SM, Shin JH, Choi HW. 2022. Availability of hydrogen peroxide solutions as a germination liquid medium for contamination-free in vitro seedling development of Cannabis sativa. Horticultural Science and Technology 40:605−13

    doi: 10.7235/HORT.20220055

    CrossRef   Google Scholar

    [44]

    Uhl L, Gerstel A, Chabalier M, Dukan S. 2015. Hydrogen peroxide induced cell death: One or two modes of action? Heliyon 1:e00049

    doi: 10.1016/j.heliyon.2015.e00049

    CrossRef   Google Scholar

    [45]

    le Roux LG, Robbertse PJ. 1994. Tuber ontogeny, morphology and vegetative reproduction of Gloriosa superba L. South African Journal Botany 60:321−24

    doi: 10.1016/S0254-6299(16)30586-5

    CrossRef   Google Scholar

    [46]

    Acemi A, Özen F. 2019. Optimization of in vitro asymbiotic seed germination protocol for Serapias vomeracea. The EuroBiotech Journal 3:143−51

    doi: 10.2478/ebtj-2019-0017

    CrossRef   Google Scholar

    [47]

    Caixeta Sousa M, Rodrigues LFOS, da Silva MB, Cruz JO, Diamante MS, et al. 2018. Productive and qualitative performance of tomato plants as a function of the application of plant growth regulators and mineral nutrients. Revista Colombiana de Ciencias Hortícolas 12:416−24

    doi: 10.17584/rcch.2018v12i2.7575

    CrossRef   Google Scholar

    [48]

    Udayan A, Kathiresan S, Arumugam M. 2018. Kinetin and Gibberellic acid (GA3) act synergistically to produce high-value polyunsaturated fatty acids in Nannochloropsis oceanica CASA CC201. Algal Research 32:182−92

    doi: 10.1016/j.algal.2018.03.007

    CrossRef   Google Scholar

    [49]

    Dalessandro G. 1973. Interaction of auxin, cytokinin, and gibberellin on cell division and xylem differentiation in cultured explants of Jerusalem artichoke1. Plant and Cell Physiology 14:1167−76

    doi: 10.1093/oxfordjournals.pcp.a074956

    CrossRef   Google Scholar

    [50]

    Saffari P, Majd A, Jonoubi P, Najafi F. 2021. Effect of treatments on seed dormancy breaking, seedling growth, and seedling antioxidant potential of Agrimonia eupatoria L. Journal of Applied Research on Medicinal and Aromatic Plants 20:100282

    doi: 10.1016/j.jarmap.2020.100282

    CrossRef   Google Scholar

    [51]

    Muhammad ZI, Maria KS, Mohammad A, Muhammad S, Zia-Ur-Rehman F, et al. 2015. Effect of mercury on seed germination and seedling growth of Mungbean (Vigna radiata (L.) Wilczek). Journal of Applied Sciences and Environmental Management 19:191−99

    doi: 10.4314/jasem.v19i2.4

    CrossRef   Google Scholar

    [52]

    Kshetrimayum E, Sahoo DP, Mitra J, Panda SK. 2017. Regulation of seed germination and the role of aquaporins under abiotic stress. International Journal of Environment, Agriculture and Biotechnology 2:238710

    doi: 10.22161/ijeab/2.2.7

    CrossRef   Google Scholar

    [53]

    Hazra A, Dasgupta N, Sengupta C, Das S. 2019. MIPS: Functional dynamics in evolutionary pathways of plant kingdom. Genomics 111:1929−45

    doi: 10.1016/j.ygeno.2019.01.004

    CrossRef   Google Scholar

    [54]

    Luo Y, Qin G, Zhang J, Liang Y, Song Y, et al. 2011. D-myo-inositol-3-phosphate affects phosphatidylinositol-mediated endomembrane function in Arabidopsis and is essential for auxin-regulated embryogenesis. The Plant Cell 23:1352−72

    doi: 10.1105/tpc.111.083337

    CrossRef   Google Scholar

    [55]

    Pathak V. 2018. Effect of starch-based hydrogel on early growth of corn. Thesis. Purdue University, US. pp. 45−53. https://docs.lib.purdue.edu/open_access_theses/1433

    [56]

    Pradhan S, Regmi T, Parmar G, Pant B. 2013. Effect of Different Media on in vitro Seed Germination and Seedling Development of Cymbidium aloifolium (L.) Sw. Nepal Journal of Science and Technology 14:51−56

    doi: 10.3126/njst.v14i1.8878

    CrossRef   Google Scholar

    [57]

    Gharari Z, Bagheri K, Karimkhanlooei G, Sharafi A. 2021. Study of tissue culture and in vitro organogenesis of Scutellaria bornmuelleri using benzylaminopurine, lsopentenyl adenine and thidiazuron. South African Journal of Botany 139:458−69

    doi: 10.1016/j.sajb.2021.03.030

    CrossRef   Google Scholar

    [58]

    Ansar A, Touqeer A, Akhtar AN, Ahmed HI. 2009. Effect of different concentrations of auxins on in vitro rooting of olive cultivar 'Moraiolo'. Pakistan Journal of Botany 41:1223−31

    Google Scholar

    [59]

    Mosoh DA, Khandel AK, Verma SK, Vendrame WA. 2024. Optimizing callus induction and indirect organogenesis in non-dormant corm explants of Gloriosa superba (L.) via media priming. Frontiers in Horticulture 3:1378098

    doi: 10.3389/fhort.2024.1378098

    CrossRef   Google Scholar

    [60]

    Mosoh DA, Khandel AK, Verma SK, Vendrame WA. 2024. Phytochemical analysis and enhanced production of alkaloids in non-dormant corm-derived callus of Gloriosa superba (L.) using plant growth regulators and abiotic elicitors. Plant Cell, Tissue and Organ Culture (PCTOC) 156:89

    doi: 10.1007/s11240-023-02674-5

    CrossRef   Google Scholar

  • Cite this article

    Mosoh DA, Khandel AK, Verma SK, Vendrame WA. 2024. Overcoming dual seed dormancy and enhancing in vitro seedling development of Gloriosa superba (L.) with a targeted sterilization approach and plant growth regulator synergy. Tropical Plants 3: e031 doi: 10.48130/tp-0024-0033
    Mosoh DA, Khandel AK, Verma SK, Vendrame WA. 2024. Overcoming dual seed dormancy and enhancing in vitro seedling development of Gloriosa superba (L.) with a targeted sterilization approach and plant growth regulator synergy. Tropical Plants 3: e031 doi: 10.48130/tp-0024-0033

Figures(12)  /  Tables(4)

Article Metrics

Article views(1898) PDF downloads(259)

ARTICLE   Open Access    

Overcoming dual seed dormancy and enhancing in vitro seedling development of Gloriosa superba (L.) with a targeted sterilization approach and plant growth regulator synergy

Tropical Plants  3 Article number: e031  (2024)  |  Cite this article

Abstract: The status of Gloriosa superba L. in the wild has been declining due to over-collection and habitat destruction. Intrinsic severe seed dormancy and low germination rates hinder the cultivation process. To conserve this plant, in vitro culture protocols have been developed to enhance seed germination and seedling growth. An effective sterilization method involving 0.15% mercuric chloride (HgCl2) for 8 min was found to eliminate contamination and yield a 100% survival rate, resulting in disinfested seeds and robust seedling growth. The most successful treatment consisted of Murashige and Skoog (MS) medium with 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP, along with 4% sucrose under a 16-h photoperiod, which achieved the highest average in vitro seed germination (9.25 out of 12 seed explants) and an impressive overall seedling survival rate of 77.08% after 30 d. Subsequent growth of two-week-old seedlings on MS medium with 1.5 mg·L−1 BAP and 1.0 mg·L−1 NAA supplemented with 30 g·L−1 sucrose and a 16-h photoperiod resulted in the greatest average seedling length (5.83 cm) and seedling root length (4.08 cm) after four weeks. Transferring the excised shoots of in vitro-grown seedlings to half-strength MS medium with 1.0 mg·L−1 IBA led to maximum root induction (84.37%) and further enhanced root development. These in vitro-grown plantlets were successfully acclimatized and transplanted under field conditions, with a 60% survival rate after 11 weeks.

    • Efficient germination is crucial for the establishment of seedlings, the growth of plants, and the preservation of species[1]. Seed moisture, humidity, and temperature are a few factors that have a big impact on germination[2]. Ecosystems are experiencing swift changes as a consequence of environmental difficulties and global warming, leading to a significant reduction in the diversity of plant species[3]. Global conservation efforts have been carried out using both in situ and ex situ methods, with seed banking being widely adopted as an effective tool to protect genetic resources[4]. Merely gathering and retaining seeds is inadequate; for effective restoration, their eventual sprouting is crucial[5]. However, many species still lack crucial knowledge about the factors that trigger dormancy release and germination, which hinders successful propagation[6]. It is essential to have a thorough understanding of seed dormancy, which is the inhibition of germination even under ideal circumstances and can result from stimuli coming from the embryo, the seed, or the fruit coat. It is crucial to overcome these obstacles to achieve germination. There are different varieties of dormancy, each requiring certain environmental circumstances to end. This knowledge is essential for the reproduction and use of plants, and it is crucial for efforts to conserve species.

      Seed germination is a complex process that begins with the absorption of water and ends with the appearance of the radicle. Plant hormones are essential for regulating seed dormancy and germination. The shift from a dormant state to an active metabolic phase is skilfully coordinated by internal gibberellins (GA), which stimulate the synthesis of hydrolytic enzymes, thus enabling the consumption of stored resources to support embryonic development[7]. Internal seed variables have an important impact on gibberellins (GA), which play a key role in seed germination[8]. Moreover, the process leading up to germination includes a substantial decrease in abscisic acid levels and a simultaneous increase in endogenous GA concentrations. The processes of imbibition and stratification facilitate this hormonal change, which is closely related to the release of dormancy[9]. Applying exogenous GA is successful in overcoming seed dormancy in certain plant species, resulting in faster seedling growth[10]. Different investigations have produced conflicting results about the presence of abscisic acid (ABA) in Gloriosa superba L. seeds. Some studies found very little or no ABA, which could mean that the seeds were physically dormant. Other studies found that chemical scarification with GA effectively overcame dormancy and sped up the germination of these seeds[10]. Nonetheless, the effectiveness of GA treatment depends on variables such as plant species, dormancy type, GA formulation, concentration, and length of treatment.

      Plant growth regulators, such as cytokinins, are pivotal in seed physiology, influencing critical processes like cell division, lateral bud expansion, and germination through their modulation of ethylene release[11]. Ethylene is a hormone that plays a critical role in seed dormancy and germination. It interacts with other plant hormones and environmental factors. It counteracts abscisic acid's (ABA) effects and promotes gibberellic acid (GA) action, influencing germination. Ethylene's effectiveness depends on GA or light, and its interaction with cytokinins like kinetin can reverse dormancy. Its impact varies depending on seed dormancy type and environmental conditions, involving complex signaling pathways and interactions with other hormones[12]. Brassinosteroids (BRs) play a crucial role in regulating plant growth and development by controlling gibberellin (GA) biosynthesis and signaling pathways[13]. They are present in various plant tissues and are involved in a range of developmental processes. The BR-regulated transcription factor BES1 binds to GA promoters, regulating their expression, which is important for seedling growth. For instance, during seed germination in cereal grains, GA signaling works together with nutrient starvation signals to stimulate hydrolases, such as α-amylase, which are essential for mobilizing nutrients[14]. In contrast, auxins accumulate in embryos to suppress germination-related genes and regulate ABA metabolism, thereby maintaining dormancy until favorable conditions prevail. Auxins like indole-3-acetic acid (IAA) are integral to both germination and root formation processes[15]. As seeds imbibe water, IAA levels rise, a precursor to root elongation, particularly promoting lateral root development, illustrating their pivotal role in seedling establishment and growth[11]. Comprehending these subtleties is essential for customized and efficient approaches to seed germination[16].

      A dormant seed is a seed that is unable to initiate germination despite favorable environmental conditions[17,18]. It can be classified into two main categories: physical dormancy, which occurs when palisade cells in the seed or fruit coat prevent water from entering, and morphological/physiological dormancy, which occurs when seeds have an embryo that is not fully developed[18,19]. These seeds require specific treatments to break dormancy and allow them to germinate[20]. Le Roux & Robbertse's 1997 study on the morphology and seed structures of Gloriosa superba L. revealed that seeds with an outer seed coat (sarcotesta) and sterilized with a 1% hypochlorite solution did not initiate germination. Some seed coats hindered water and oxygen absorption, causing delays[21]. However, removing the sarcotesta enhanced germination, which began 13 d after removing the sarcotesta at room temperature but with mixed results, especially at different temperatures, as germination remained unpredictable due to physiological variations among seeds. Researchers have looked into various pre-sowing techniques to enhance seed germination; however, the results remain poor[22,23].

      Surface sterilization of seeds and explants is necessary to get rid of any microbes that could be harmful before in vitro regeneration techniques are used[24]. Efficiency is crucial in biotechnology-driven breeding, as it seeks to expedite the cultivation of superior plants within a limited time frame[25]. Nevertheless, the challenge of attaining efficient sterilization while maintaining plant development is a multifaceted undertaking[26]. The kind, size, and age of the explant, as well as the growing conditions, all have an impact on the effectiveness of disinfection[27]. Various disinfectants, including sodium hypochlorite (NaClO), hydrogen peroxide (H2O2), mercury II chloride (HgCl2), and others can be used depending on the degree of contamination and prevailing conditions. Extended duration and increased doses of disinfectants typically yield improved asepsis outcomes, but this may affect the viability of explants[28]. To get effective sterilization in vitro, it is important to vary the exposure time and disinfectant concentration based on the unique properties of the explants and species that were chosen.

      The rising global demand for Gloriosa superba L. in export markets exacerbates its depletion and heightens the risk of extinction[29,30]. Historically, unsustainable harvesting practices have predominantly targeted tuber extraction, driven by the therapeutic benefits associated with this plant. Although countries such as India and Zimbabwe have enacted protective measures to regulate wild resource collection, illegal harvesting persists due to scarcity and escalating market demand[31]. Moreover, Gloriosa superba L. encounters challenges such as low, unpredictable, and insufficient seed germination rates, as well as susceptibility to pests in its natural habitat[32]. Meeting commercial demands requires large-scale cultivation. Although vegetative propagation is common in horticulture, it is hindered by slow growth and limited multiplication rates, which prolong the reproductive cycle. In vitro cultivation of Gloriosa superba L. seeds provides a solution to improve germination rates and accelerate growth. Seeds are preferred over vegetative parts for conservation purposes as they preserve broader genetic diversity[33].

      A recent study used BET surface area analysis and 3D X-ray micro-tomography to investigate the seed coat structure of Gloriosa superba L. The study revealed that the sarcotesta and endosperm exhibit minimal porosity, rendering them impervious to water during germination[22]. Traditional techniques such as acid or mechanical scarification and water soaking are commonly used to facilitate water and oxygen absorption in hard-coated seeds, hence encouraging germination. However, despite these methods, germination rates remain low and inconsistent[22]. In vitro propagation is a versatile technique widely used in applied plant research for many purposes, such as preserving germplasm, conserving species, propagating clones on a large scale, and addressing limitations in traditional propagation methods[33,34]. It is highly beneficial for boosting seed germination by circumventing dormancy and optimizing growth conditions, resulting in reduced germination time and improved shoot and root development.

      Numerous studies have researched the sterilization and germination processes of Gloriosa superba L. seeds, consistently reporting low germination rates[35]. As previously mentioned, optimizing sterilization techniques can greatly improve seed germination rates[33]. Therefore, establishing reliable protocols for sterilizing Gloriosa superba L. seeds and utilizing them for in vitro micropropagation is essential. This effort is critical for conserving and sustainably utilizing this species, particularly due to its vulnerable status in its natural habitat.

      This study focuses on the crucial matter of seed dormancy in Gloriosa superba L. Although the exact dormancy classification of this species is unknown, a dual dormancy model is proposed, which includes both physical and physiological components. Through using the right seed sterilization method along with the right mix of gibberellic acid (GA3) and other plant growth regulators, this study suggests that it is possible to break dormancy and improve seed germination and seedling growth. This study seeks to develop a reliable method for sterilizing Gloriosa superba L. seeds and refine a methodology for in vitro propagation to efficiently produce Gloriosa superba L. plantlets from seeds. The findings of this research will significantly contribute to conservation efforts.

    • Fresh seeds of Gloriosa superba L. were carefully collected from mature and strong plants that were thriving in the Pachmarhi Biosphere Reserve (PBR) located in Madhya Pradesh, India (Fig. 1). The harvested seeds were sorted into sets of 100 and weighed accurately. These seeds were then stored in specialized containers under ambient conditions (temperature and relative humidity of 25 ± 2 °C and 50%, respectively), in preparation for further analysis. The culture medium used in this study was Murashige and Skoog (MS) medium, which was purchased from Merck (Mumbai, India). Additionally, Gibberellic acid (GA3), 6-benzylaminopurine (BAP), Indole-3-acetic acid (IAA), Indole-3-butyric acid (IBA), Kinetin (KN), 1-Naphthalene acetic acid (NAA), N6-(2-Isopentenyl) adenine (2-iP), and sucrose were sourced from Sigma-Aldrich (Bengaluru, India). Sterilizing agents such as mercuric chloride (HgCl2), sodium hypochlorite (NaClO), and hydrogen peroxide (H2O2) were obtained from Merck (Mumbai, India).

      Figure 1. 

      Gloriosa superba L. seeds were obtained from the Pachmarhi Biosphere Reserve (Madhya Pradesh, India) and cultured using in vitro techniques. (a) Batch of pre-sterilized seeds; (b) Seed sterilization process taking place in laminar air flow (LAF); (c) Seed germination; (d) Healthy in vitro seedling germination. Scale bar = 2 cm.

    • The seeds of Gloriosa superba L. were surface sterilized using various concentrations (w/v) and exposure times (minutes) of three sterilants: HgCl2 (Merck), NaClO (Merck), and H2O2 (Merck) (Table 1). The seeds were first washed for 15−20 min under running tap water in conjunction with a sifting sieve to remove dirt and pulp. Subsequently, they underwent treatment with a 5% (v/v) detergent solution (Teepol), vortexed for 8 min, followed by five rinses with tap water. After that, they were immersed in a 2% (w/v) Bavistin fungicide solution, vortexed for 10 min, and then rinsed five times with tap water. Pre-sterilized seeds were transferred to a controlled environment (Laminar Air Flow (LAF)) following appropriate sterilization procedures for both the workstation and hands and washed five times with sterile distilled water to remove any traces of fungicide. The seeds were then briefly submerged in 70% (v/v) ethanol for 20 s, rinsed five times with sterile distilled water, and subjected to sterilization agents with varying concentrations (0.05%, 0.1%, 0.15%) and exposure times (2, 5, and 8 min). Afterward, the seeds were rinsed five times with sterile distilled water for 8 min to remove any residual sterilizing agents. The sterilized seeds were then ready for inoculation into the culture medium.

      Table 1.  Concentrations and exposure durations of mercuric chloride (HgCl2), sodium hypochlorite (NaClO), and hydrogen peroxide (H2O2) used to assess the contamination levels and survival rate of in vitro germinated Gloriosa superba L. seedlings.

      Treatments Sterilant Concentration (w/v) Exposure time (min)
      T1 HgCl2 0.05% 2
      T2 0.05% 5
      T3 0.05% 8
      T4 0.1% 2
      T5 0.1% 5
      T6 0.1% 8
      T7 0.15% 2
      T8 0.15% 5
      T9 0.15% 8
      T10 NaClO 0.5% 2
      T11 0.5% 5
      T12 0.5% 8
      T13 1.0% 2
      T14 1.0% 5
      T15 1.0% 8
      T16 1.5% 2
      T17 1.5% 5
      T18 1.5% 8
      T19 H2O2 5.0% 2
      T20 5.0% 5
      T21 5.0% 8
      T22 7.5% 2
      T23 7.5% 5
      T24 7.5% 8
      T25 10% 2
      T26 10% 5
      T27 10% 8

      The seeds were inspected after sterilization to ensure that they were free from unwanted after-effects of the treatment. They were then cultured in flasks containing half-strength MS medium supplemented with 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP. The pH of the medium was set to 5.8, and it was solidified using 0.8% agar (Merck, India). Each replicate consisted of eight seeds cultivated in separate 250-mL flasks, with four replications per treatment. The cultures were maintained under a 16-h photoperiod with an 80 μmol·m−2·s−1 photosynthetic photon flux density provided by white fluorescent tubes (40 W; Philips, India). The temperature and relative humidity were set at 25 ± 2 °C and 60%, respectively. After 4 weeks, the effectiveness of different sterilization agents, concentrations, and immersion times on contamination and survival were determined (Fig. 1).

    • The Gloriosa superba L. seeds were cleaned with the best sterilization method from the previous experiment. Then, the sterilized seeds were placed in 250-mL conical flasks containing 50 ml of full-strength MS medium supplemented with 30 g·L−1 sucrose and different concentrations of PGRs (GA3, BAP, and NAA) to evaluate their effects on in vitro seed germination (Table 2). The pH of the medium was adjusted to 5.8, and 0.8% agar (HiMedia, India) was added to solidify the medium. The flasks were sealed with non-absorbent cotton plugs and autoclaved at 121 °C for 20 min under 104 kPa pressure. Four replications were used per treatment; each replicate contained 12 seeds cultured in an individual 250-mL flask.

      Table 2.  Various concentrations and combinations of Gibberellic acid (GA3), 6-benzylaminopurine (BAP), Kinetin (KN), and 1-naphthalene acetic acid (NAA) used to evaluate in vitro germination of Gloriosa superba L. seeds.

      Treatment PGRs (mg L−1)
      GA3 BAP
      T1 0.2 0.2
      T2 0.5 0.5
      T3 1.0 1.0
      T4 1.5 1.5
      T5 2.0 2.0
      T6 2.5 2.5
      GA3 KN
      T7 0.2 0.2
      T8 0.5 0.5
      T9 1.0 1.0
      T10 1.5 1.5
      T11 2.0 2.0
      T12 2.5 2.5
      GA3 NAA
      T13 0.2 0.2
      T14 0.5 0.5
      T15 1.0 1.0
      T16 1.5 1.5
      T17 2.0 2.0
      T18 2.5 2.5
      T19 (Control) 0 0

      All the cultures were kept under a 10/14-h light/dark photoperiod using white fluorescent tubes (40 W; Philips, India) that provided 80 μmol·m−2·s−1 photosynthetic photon flux density. The greenhouse's relative humidity was 60%, and the temperature was maintained at 25 ± 2 °C. After 3, 5, 9, 13, 21, and 30 d, the number of germinated seeds and seedling survival were counted to assess seed germination.

      In this study, seed germination was primarily defined by the emergence of a sprout, typically measuring 3 cm in length. However, regardless of the sprout's length, the appearance of specific morphological features such as the plumular leaves, cotyledonary sheath, swollen stem base, and primary root confirmed germination, accounting for variations in treatment conditions and seed health. The germination experiment was conducted for 30 d, with all seeds monitored for these key developmental markers throughout the entire duration.

    • Following two weeks of seed germination on a full-strength MS medium containing 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP, fresh seedlings were produced. The seedlings were transferred to new, full-strength MS media containing 30 g·L−1 sucrose and varying amounts of BAP, NAA, 2-iP, and IAA. The different treatment groups were used to evaluate their effects on the morphological characteristics of the seedlings (Table 3). The pH of these media was maintained at 5.8, and all the cultures were kept under a 16-h photoperiod with a photosynthetic photon flux density of 80 μmol·m−2·s−1 given by white fluorescent tubes (40 W; Philips, India). The ambient relative humidity and temperature were 60% and 25 ± 2 °C, respectively.

      Table 3.  Various concentrations and combinations of 6-benzylaminopurine (BAP), 1-naphthalene acetic acid (NAA), N6-(2-Isopentenyl) adenine (2-iP), and Indole-3-acetic acid (IAA) used to evaluate in vitro growth and development Gloriosa superba L. seedlings.

      Treatments PGRs (mg·L−1)
      BAP NAA
      T1 0.5 0.2
      T2 1.0 0.5
      T3 1.5 1.0
      2ip IAA
      T4 0.5 0.2
      T5 1.0 0.5
      T6 1.5 1.0
      T7 (control) 0 0

      All parameters, including shoot length, root length, root/shoot ratio, seedling length, and fresh weight, were measured twice. This was done before and after a 28-d seedling enhancement experiment. The only exception was the measurement of seedling biomass dry weight, which was taken once at the end of the experiment. To measure the fresh and dry weight parameters of the seedlings, an electronic balance with a precision of 0.01 mg was used. Fresh weight was measured directly (under aseptic conditions before this experiment), whereas, for dry weight measurement, seedlings were subjected to a two-step drying process. First, they were dried in an oven at 105 °C for 30 min, and then at 75 °C until a steady weight was achieved.

    • Microshoots were cut from mature seedlings and placed in culture media (half-strength MS basal media) with varying concentrations of IBA (0.5, 1.0, 1.5 mg·L−1), IAA (0.5, 1.0, 1.5 mg·L−1), and NAA (0.5, 1.0, 1.5 mg·L−1), as shown in Table 4. The pH of the half-strength MS basal media was adjusted to 5.8, and 0.8% (w/v) agar (HiMedia, India) was added to solidify the medium.

      Table 4.  Various concentrations of Indole-3-butyric acid (IBA), Indole-3-acetic acid (IAA), and 1-naphthalene acetic acid (NAA) used to assess in vitro root development in young shoots excised from Gloriosa superba L. seedlings.

      Treatments 1/2 MS + auxins (mg·L−1)
      IBA
      T1 0.5
      T2 1.0
      T3 1.5
      IAA
      T4 0.5
      T5 1.0
      T6 1.5
      NAA
      T7 0.5
      T8 1.0
      T9 1.5
      T10 (control) 0.0

      The flasks with the different treatments of media were sealed with non-absorbent cotton plugs and autoclaved at 121 °C for 20 min at 104 kPa of pressure. This was done before the microshoot culture. Each treatment was replicated four times, with each replicate consisting of eight axenic microshoots cultivated in separate 250-mL flasks. Culture conditions were maintained as per the previously provided description. The cultures were checked after six weeks, and information was gathered on how many microshoots grew roots, how quickly they responded to the rooting treatment, how long it took for roots to form on each microshoot, and how long the roots were in centimeters.

    • Before placing plantlets in small polyethylene bags, trays, pots, or thermocol cups (7 cm in diameter) filled with sterilized vermiculite and soil (1:1), they were rinsed with deionized water to eliminate surplus media. During the initial stage of acclimatization, the plantlets were kept under a 16-h photoperiod. They were exposed to white fluorescent tubes (40 W; Philips, India) that emitted a photosynthetic photon flux density of 50 μmol m−2 s−1. The plantlets were enclosed in polyethylene bags with small air holes to ensure high relative humidity (RH) of 90% and to prevent dehydration. The temperature in the culture room (CR) was maintained at 25 ± 2 °C. The polythene casings were regularly removed for 1 h. For two weeks, all potted plantlets were watered with 10 ml of a half-strength Murashige and Skoog (MS) basal salt solution (without sucrose and myo-inositol) every 4 d.

      In the second stage, which occurred during the third week, the plantlets were transferred into medium-sized polyethylene bags, plastic cups, or thermocol cups filled with a mixture of garden soil, sand, and vermiculite in a ratio of 2:1:1 (volume to volume). The plantlets were housed in a shade net house (USNH) for 1 week, receiving regular mistings of tap water. The relative humidity (RH) was gradually decreased by 50%, and the plantlets were thereafter transplanted directly to the experimental field and home garden for a duration of 11 weeks to facilitate their continued growth and development.

      When plantlets were moved, they were put in three different conditions: first, they were put in a controlled culture room for 2 weeks in a mixture of sterilized vermiculite and soil (1:1); then, they were put in a net house for 1 week in a mixture of garden soil, sand, and vermiculite (2:1:1) in the shade (USNH); finally, they were put out in the field for 11 weeks in direct sunlight (DSL). Information was gathered for a maximum duration of 14 weeks after the initiation of microplantlet acclimation. After transplanting the microplantlets onto the stated potting mixtures, weekly observations were conducted. The overall count of microplantlets recorded per treatment was 56 (with four replicates, each containing 14 microplantlets). The survival percentage of the regenerated plantlets was calculated using the formula:

      Survivalrate(%)=No.ofsurvivingregeneratedplantsTotalno.oftransplantedregeneratedplants×100%

      The data represents the mean and standard error (SE).

    • A fully randomized experimental design was utilized for all trials, wherein seeds and seedlings were randomly chosen and organized into groups for each treatment. The seed sterilization, seed germination, seedling development, and rooting studies were conducted using four replicates for each treatment level. Each replication consisted of eight seeds, 12 seeds, 12 2-week-old seedlings, and eight microshoots, respectively. The experiments were performed twice.

      After 4 weeks, data was collected for all parameters in each experiment. The percentage response to treatment was determined by dividing the total number of explants or microshoots that exhibited a response by the total number of replicates and then multiplying the result by 100. The seed germination percentage was determined by dividing the number of healthy seedlings by the total number of replicates and then multiplying the result by 100. The seed contamination rate was determined by dividing the total number of contaminated seeds by the total number of replicates and then multiplying the result by 100. The seedling survival rate was determined by dividing the total number of surviving seedlings by the total number of replicates and then multiplying the result by 100.

      The determination of normality was conducted using the Shapiro-Wilk test. If the normality test had a p-value greater than or equal to 0.05, a parametric test, specifically a one-way ANOVA at a significance level of 0.05, was employed to compare the means. Conversely, if the p-value was less than or equal to 0.05, a non-parametric test, specifically a Kruskal-Wallis test at a significance level of 0.05, was used to compare the means. The data were gathered and subjected to one-way analysis of variance (ANOVA) and/or the Kruskal-Wallis test using R software (version 4.4.0). The mean separation was performed using Tukey's honestly significant difference (HSD) test with a significance level (α) of 0.05. The data were shown as mean values with a standard error. Different letters in the figures indicated significant differences at a significance level of p < 0.05.

    • The highest contamination frequency (68.75%) and lowest contamination frequency (0.00%) were found at 5% H2O2 for 2 min and 10% H2O2 for 5 min, and 8 min, respectively, which were significantly different (p < 0.001) (Fig. 2a, b). Furthermore, there were significant differences in explant viability between treatments (p < 0.001) (Fig. 2c, d). Explant viability was highest (96.88%) in 7.5% H2O2 for 8 min of immersion.

      Figure 2. 

      Effects of various concentrations of hydrogen peroxide (H2O2) and immersion times on Gloriosa superba L. seed sterilization after 4 weeks of culture. (a) Mean seed contamination, (b) seedling contamination rate, (c) mean number of seedlings that survived, and (d) seedling survival rate are shown. Treatments are T19: 5.0% H2O2 for 2 min; T20: 5.0% H2O2 for 5 min; T21: 5.0% H2O2 for 8 min; T22: 7.5% H2O2 for 2 min; T23: 7.5% H2O2 for 5 min; T24: 7.5% H2O2 for 8 min; T25: 10% H2O2 for 2 min; T26: 10% H2O2 for 5 min; T27: 10% H2O2 for 8 min. Bars indicate mean ± SE. Different letters indicate significant difference by Tukey's test at p ≤ 0.05.

    • The different treatments demonstrated a significant difference in contamination and explant viability (p < 0.001). The highest contamination was recorded for 0.05% HgCl2 for 2 min of immersion, while no contamination was recorded for 0.15% HgCl2 for 8 min of immersion (Fig. 3a, b). The treatment with 0.15% HgCl2 for 8 min of immersion had the highest explant viability (100.00%). However, the lowest explant viability (21.87%) was observed after exposure to 0.05% HgCl2 for 2 min (p < 0.001) (Fig. 3c, d).

      Figure 3. 

      Effects of various concentrations of mercuric chloride (HgCl2) and immersion times on Gloriosa superba L. seed sterilization after 4 weeks of culture. (a) Mean seed contamination, (b) seedling contamination rate, (c) mean number of seedlings that survived, and (d) seedling survival rate are shown. Treatments are T1: 0.05% HgCl2 for 2 min; T2: 0.05% HgCl2 for 5 min; T3: 0.05% HgCl2 for 8 min; T4: 0.1% HgCl2 for 2 min; T5: 0.1% HgCl2 for 5 min; T6: 0.1% HgCl2 for 8 min; T7: 0.15% HgCl2 for 2 min; T8: 0.15% HgCl2 for 5 min; T9: 0.15% HgCl2 for 8 min. Bars indicate mean ± SE. Different letters indicate significant difference by Tukey's test at p ≤ 0.05.

    • The results showed that 0.5% sodium hypochlorite for 2 min of immersion and 1.5% sodium hypochlorite for 8 min of immersion resulted in the highest (84.38%) and lowest (6.25%) contaminations, respectively (p < 0.001) (Fig. 4a, b). Also, the explant viability was highest (93.75%) when it was immersed in 1.5% sodium hypochlorite for 8 min. This was statistically significantly different from the lowest explant viability (15.63%) (p < 0.001) (Fig. 4c, d).

      Figure 4. 

      Effects of different sodium hypochlorite (NaClO) concentrations and immersion times on Gloriosa superba L. seed sterilization after 4 weeks of culture. (a) Mean seed contamination, (b) seedling contamination rate, (c) mean number of seedlings that survived, and (d) seedling survival rate. Treatments are T10: 0.5% NaClO for 2 min; T11: 0.5% NaClO for 5 min; T12: 0.5% NaClO for 8 min; T13: 1.0% NaClO for 2 min; T14: 1.0% NaClO for 5 min; T15: 1.0% NaClO for 8 min; T16: 1.5% NaClO for 2 min; T17: 1.5% NaClO for 5 min; T18: 1.5% NaClO for 8 min. Bars indicate mean ± SE. Different letters indicate significant difference by Tukey's test at p ≤ 0.05.

    • For 30 d, Gloriosa superba L. seeds showed increased germinability in response to the various treatments. Except for the 3rd day (p < 0.4557), the different treatments showed a significant difference only after the 5th day of germination (p < 0.001) (Fig. 5ag). On the 3rd, 5th, 9th, 13th, 21st, and 30th days, treatment with 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP resulted in the highest average seed germination of 0.25, 3.00, 5.75, 7.25, 9.25, and 9.25, respectively (Figs 6, 7). Treatment with 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP showed the highest seedling survival rate (77.08%), while treatment with 0.2 mg·L−1 GA3 and 0.2 mg·L−1 NAA showed the lowest (50.00%).

      Figure 5. 

      Effect of plant growth regulators (PGRs) on Gloriosa superba L. seed germination after 30 d of culture. (a) Mean seed germination after 3 d, (b) mean seed germination after 5 d, (c) mean seed germination after 9 d, (d) mean seed germination after 13 d, (e) mean seed germination after 21 d, (f) mean seed germination after 30 d, and (g) seedling survival rate after 30 d are shown. Treatments are T1: 0.2 mg·L−1 GA3 (Gibberellic acid), 0.2 mg·L−1 BAP (6-benzylaminopurine); T2: 0.5 mg·L−1 GA3, 0.5 mg·L−1 BAP; T3: 1.0 mg·L−1 GA3, 1.0 mg·L−1 BAP; T4: 1.5 mg·L−1 GA3, 1.5 mg·L−1 BAP; T5: 2.0 mg·L−1 GA3, 2.0 mg·L−1 BAP; T6: 2.5 mg·L−1 GA3, 2.5 mg·L−1 BAP; T7: 0.2 mg·L−1 GA3, 0.2 mg·L−1 KN (Kinetin); T8: 0.5 mg·L−1 GA3, 0.5 mg·L−1 KN; T9: 1.0 mg·L−1 GA3, 1.0 mg·L−1 KN; T10: 1.5 mg·L−1 GA3, 1.5 mg·L−1 KN; T11: 2.0 mg·L−1 GA3, 2.0 mg·L−1 KN; T12: 2.5 mg·L−1 GA3, 2.5 mg·L−1 KN; T13: 0.2 mg·L−1 GA3, 0.2 mg·L−1 NAA (1-Naphthalene acetic acid); T14: 0.5 mg·L−1 GA3, 0.5 mg·L−1 NAA; T15: 1.0 mg·L−1 GA3, 1.0 mg·L−1 NAA; T16: 1.5 mg·L−1 GA3, 1.5 mg·L−1 NAA; T17: 2.0 mg·L−1 GA3, 2.0 mg·L−1 NAA; T18: 2.5 mg·L− 1 GA3, 2.5 mg·L−1 NAA; T19: Control (media without PGRs). Bars indicate mean ± SE. Different letters indicate significant difference by Tukey's test at p ≤ 0.05.

      Figure 6. 

      Gloriosa superba L. seed germination. (a) & (b) Imbibition, seed swelling, and protrusion of the radicle. (c) & (d) Seed and seedling germination. Scale bar = 2 cm.

      Figure 7. 

      Effects of various concentrations of PGRs (GA3, BAP, and NAA) on Gloriosa superba L. seed germination in full-strength MS medium supplemented with 30 g·L−1 sucrose after 30 d of culture. (a) & (b) Initial seedling developmental stage on 1.5 mg·L−1 GA3 (Gibberellic acid) + 1.5 mg·L−1 NAA (1-Naphthalene acetic acid) fortified medium. (c) & (d) Initial seedling developmental stage on 1.5 mg·L−1 GA3 + 1.5 mg·L−1 KN (Kinetin) fortified medium. (e) & (f) Initial seedling developmental stage on 1.5 mg·L−1 GA3 + 1.5 mg·L−1 BAP fortified medium. Scale bar = 2 cm.

    • Except for the shoot-to-root ratio (p < 0.515), the different PGR concentrations resulted in significant differences in seedling shoot length (p < 0.0155), seedling length (p < 0.0034), seedling root length (p < 0.00228), fresh weight (p < 0.001), and dry weight (p < 0.001) (Fig. 8af). The treatment of seedlings with 1.5 mg·L−1 BAP and 1.0 mg·L−1 NAA showed the highest average seedling shoot length (4.2 cm), seedling length (5.83 cm), seedling root length (4.08 cm), fresh weight (334 mg), and dry weight (39.1 mg) except for the shoot-to-root ratio, which showed no significant difference (p < 0.515) (Fig. 9).

      Figure 8. 

      Effect of plant growth regulators (PGRs) on Gloriosa superba L. seedling morphological traits after four weeks of culture. (a) Mean seedling shoot length, (b) mean seedling length, (c) mean seedling shoot-to-root ratio, (d) mean seedling root length, (e) mean seedling fresh weight, and (f) mean seedling dry weight are shown. Treatments are T1: 0.5 mg·L−1 BAP (6-benzylaminopurine), 0.2 mg·L−1 NAA (1-Naphthalene acetic acid); T2: 1.0 mg·L−1 BAP, 0.5 mg·L−1 NAA; T3: 1.5 mg·L−1 BAP, 1.0 mg·L−1 NAA; T4: 0.5 mg·L−1 2iP (N6-(2-isopentenyl) adenine), 0.2 mg·L−1 IAA (Indole 3-acetic acid); T5: 1.0 mg·L−1 2iP, 0.5 mg·L−1 IAA; T6: 1.5 mg·L−1 2iP, 1.0 mg·L−1 IAA; T7: control (media without PGRs). Bars indicate mean ± SE. Different letters indicate significant difference by Tukey's test at p ≤ 0.05.

      Figure 9. 

      Seedlings of Gloriosa superba L. cultured in shooting and seedling enhancement media to test the effects of different plant growth regulator combinations on the morphological traits and biomass of seedlings. (a) Germinated seeds before transfer into seedling enhancement media. (b) Initiation of seedling development on 1.5 mg·L−1 BAP (6-benzylaminopurine) + 1.0 mg·L−1 NAA (1-Naphthalene acetic acid). (c)−(d) Seedling growth on 1.5 mg·L−1 BAP + 1.0 mg·L−1 NAA after 4 weeks. Scale bar = 2 cm.

      Treatment with 1.0 mg·L−1 2iP and 0.5 mg L−1 IAA produced the second-best results for increasing average seedling length (4.71 cm) (Fig. 8b), while treatment with 1.0 mg·L−1 BAP and 0.5 mg·L−1 NAA produced the second-best results for increasing average seedling shoot length (3.51 cm), seedling root length (3.90 cm), fresh weight (294 mg), and dry weight (32 mg). Both treatments (1.0 mg·L−1 BAP, 0.5 mg·L−1 NAA, and 0.5 mg·L−1 2iP, 0.2 mg·L−1 IAA) produced the highest average shoot-to-root ratio (1.11) (Fig. 8c).

    • Young seedlings of Gloriosa superba L. transferred from the seedling enhancement treatments were cultured in half-strength MS media supplemented with various concentrations of IBA (0.5–1.5 mg·L−1), IAA (0.5–1.5 mg·L−1) and NAA (0.5–1.5 mg·L−1) for rooting (Fig. 10). The responses to root formation changed depending on the type and amount of auxin added to the medium. These changes were significant (p < 0.001), but the average root length stayed the same. The best treatment for rooting was 1.0 mg·L−1 IBA, with a rooting rate of 84.37% (Fig. 10b). 1.0 mg·L−1 IBA induced healthy adventitious roots; the rooting speed was the fastest (within 7 d) (Fig. 10c), and no calluses appeared at the bases of the adventitious roots.

      Figure 10. 

      Effect of PGRs on Gloriosa superba L in vitro morphogenetic response to root induction from seedling derived microshoot explants. (a) Mean microshoots forming roots, (b) response rate to rooting treatment, (c) mean days required for root induction, (d) mean root length, and (e) mean root per explant. Treatments are T1: 0.5 mg·L−1 IBA (Indole-3-butyric acid); T2: 1.0 mg·L−1 IBA; T3: 1.5 mg·L−1 IBA; T4: 0.5 mg·L−1 IAA (Indole 3-acetic acid); T5: 1.0 mg·L−1 IAA; T6: 1.5 mg·L−1 IAA; T7: 0.5 mg·L−1 NAA (1-Naphthalene acetic acid); T8: 1.0 mg·L−1 NAA; T9: 1.5 mg·L−1 NAA; T10: Control (media without PGRs). Bars indicate mean ± SE. Different letters indicate significant difference by Tukey's test at p ≤ 0.05.

    • Plantlets transferred ex vitro on a substrate of sterilized vermiculite-soil 1:1 (v/v) and grown in a culture room for 14 d, followed by 7 d under shade in a net house on a substrate of garden soil-sand-vermiculite 2:1:1 (v/v), were readily acclimatized at a rate of 100% (Fig. 11a, 12ae). During the next 11 weeks of growth in the field under direct sunlight, the survival rate dropped to 60% (Fig. 11a, 12f, g). All of the surviving plants flowered, produced microtubers, grew to normal plant height, and had multiple healthy leaves (p < 0.001) (Figs 11be; 12hj).

      Figure 11. 

      Acclimatization of in vitro regenerated seedlings of Gloriosa superba L. (a) Plant survival, (b) plant height in cm, (c) number of leaves per plant, (d) number of flowers per plant, and (e) number of microtubers per plant, measured after 2 weeks of transplant in sterilised (vermiculite + soil, 1:1) grown in culture room (CR); 1 week of transplant in garden soil + sand + vermiculite, 2:1:1 under shade in net house (USNH), and finally, 11 weeks of transplant in the field under direct sun light (DSL).

      Figure 12. 

      Acclimatization of in vitro regenerated seedlings of Gloriosa superba L. (a) Microshoots transplanted into sterilized substrate (vermiculite + soil, 1:1) grown in culture room (CR). (b) Plantlets in chill trays containing a mixture of garden soil + sand + vermiculite, 2:1:1 under shade in net house (USNH). (c) Direct transplantation in the field under direct sun light (DSL). (d) Fully grown plant with developed flowers at the later stage of acclimatization in the field under direct sun light (DSL). (e) Some microtubers harvested at the later stage of acclimatization. Scale bar = 2 cm.

    • This work focused on enhancing the in vitro germination of Gloriosa superba L. seeds, aiming to resolve the problem of seed dormancy and sluggish and inconsistent seed germination commonly encountered in natural and traditional cultivation approaches. The use of an effective seed sterilization method, combined with the strategic application of GA3 and other plant growth regulators, resolved seed dormancy and resulted in significant improvements in both seed germination and seedling growth, thus demonstrating the superiority of in vitro plant tissue culture techniques in micropropagation via seed explants and conservation.

    • The presence of microorganisms, including bacteria, yeast, and fungi, is the primary source of losses in plant in vitro cultivation due to contamination. These microorganisms engage in a competition for resources with plant tissues, which often leads to a higher rate of mortality in the culture. Nevertheless, their existence can also lead to inconsistent growth, tissue death, decreased shoot multiplication, and diminished root development. Surface sterilization of explants can be challenging, and even small errors during the sterilization procedure can lead to a waste of time, effort, and resources. This can have significant financial repercussions if not addressed effectively.

      Furthermore, living materials must retain their biological activity while undergoing sterilization, with only contaminants being destroyed. Therefore, explants are subjected to surface sterilization using sterilants at suitable concentrations for a given duration. Sen et al. discovered that the negative effects on germination (%) and plantlet development became more noticeable after an 8-min treatment. This suggests that growth is significantly slowed after this time. Therefore, a standardized immersion period limit of 8 min was implemented for all seed sterilization trials[36].

      Sodium hypochlorite, mercuric chloride, and hydrogen peroxide are frequently employed as sterilants to achieve surface sterilization of plant and seed material from different species[37]. Nevertheless, these compounds sometimes prove inadequate for successfully eliminating contaminants, particularly when it comes to seeds that have been harvested from an open field and stored under unsterile conditions. The current study aimed to determine the optimal sterilization protocol for Gloriosa superba L. seeds by testing various concentrations of mercuric chloride (0.05%, 0.1%, and 0.15%), sodium hypochlorite (0.5%, 1.0%, and 1.5%), and hydrogen peroxide (5.0%, 7.5%, and 10%) at different immersion times (2, 5, and 8 min).

      Mercuric chloride exhibits potent antibacterial properties, effectively eliminating both fungi and bacteria. However, it can also be detrimental to seeds and plant materials, perhaps causing their demise. It is potentially the most efficient agent for sterilizing soil-borne and epiphytic fungi in seeds. It is favored over NaClO and H2O2 due to its higher level of activity. Mercuric chloride is extremely poisonous, but sterilants like hypochlorite are consistently safer, even when used in high quantities[38]. Unlike earlier studies that used a 0.1% concentration of HgCl2, this one showed that the seeds were not sterilized even when they were exposed to this concentration and for as long as they were submerged[39]. Instead, the best benefits came from utilizing a slightly higher concentration (0.15%) and the longest possible immersion time (8 min). This confirmed that the tissue samples were fully functional and free from any contamination. This highlights the possibility of enhancing the effectiveness of sterilization by raising both the concentration and duration of immersion. Gloriosa superba L. seeds had a strong sarcotesta that protected them from the possible harmful effects of high HgCl2 concentrations and immersion durations.

      The use of hypochlorite salts for disinfection can be traced back to the mid-18th century[40]. Most disinfection products primarily consist of chlorine-based industrial solutions[41]. Of all the NaClO concentrations that were tested, the best results were seen when the highest concentration (1.5%) was combined with the longest immersion time (8 min). This led to the highest percentage of seedling survival (93.75%) and the lowest percentage of contamination (6.25%). In contrast to HgCl2, where phytotoxicity had a direct effect on the viability of explants, the success of sterilization had a bigger effect on the survival of seedlings in NaClO. Significantly, seedling survival was negatively impacted when the immersion time with NaClO was less than 5 min, irrespective of concentration. This emphasizes the crucial importance of the duration of immersion. This discovery is consistent with prior studies on Ficus religiosa seeds, which showed that higher concentrations of NaClO and longer periods of immersion had a substantial positive effect on sterilization[42]. Similarly, a study conducted on Achyranthes aspera seeds found that higher doses of NaClO were effective in sterilizing the seeds[36].

      Hydrogen peroxide is a widely recognized compound that exhibits strong oxidizing properties[43]. According to the study's findings, placing seeds in a 7.5% H2O2 solution for 8 min led to the highest rate of seedling survival (96.88%) and a contamination rate of 3.13%. On the other hand, exposing the seeds to a greater concentration of H2O2 (10%) for a minimum of 5 min resulted in complete sterilization, but it led to decreased seedling survival. This suggests that the concentration of H2O2 affects the effectiveness of sterilization, with higher amounts having a detrimental effect on the viability of the explant. The results align with earlier studies that have shown that greater concentrations of H2O2 can decrease contamination but may also harm the viability of the explants[44]. While several studies indicate that sodium hypochlorite is more efficacious in contamination control, alternative viewpoints contend that hydrogen peroxide may be preferable, leading to reduced contamination levels and increased germination rates[37]. Data showing that H2O2 is a potent sterilizing agent and also aids seed germination by suppressing germination inhibitors support this viewpoint[43]. This phenomenon could perhaps elucidate the reason behind the unusually high survival of the explants, even when exposed to higher concentrations (up to 10%) and longer immersion durations (up to 8 min), without any evidence of contamination.

    • Gloriosa superba L. seeds are pale orange, sarcotestal, and desiccated, with a dormancy phase lasting approximately four months. Under optimal conditions, mature seeds initiate germination approximately 2 weeks following water absorption[45]. The linear embryo is 2.0–2.5 mm in length and is enclosed within the mostly proteinaceous endosperm. During germination, the haustorial cotyledon undergoes elongation, while the embryo axis is expelled from the seed[46] (Fig. 6). Hypogeal germination is characterized by the elongation of the stem axis, keeping the cotyledon below the ground. Root primordia originate internally within the stem tissue, resulting in the development of a widespread adventitious root system. The cotyledon gives rise to a well-developed vascular system that extends into the primary root and the initial leaf primordia responsible for photosynthesis (Fig. 6). Once the seedling has grown a minimum of two photosynthetic leaves, the lower part of the stem begins to enlarge. The seedling undergoes a rosette stage, characterized by the presence of two or more leaves, prior to the elongation of the internode below the apical bud. The swelling stem area predominantly comprises two buds, occasionally three or four, depending on the number of leaves in the rosette stage. The tuberous hypopodium initially extends horizontally, then responds to gravity and transforms into a bloated, cylindrical subterranean tuberous structure with a vertical orientation. An 'anatropous bud' is formed when the basal part of the bud is pushed towards the terminal position due to hyponastic development[45]. Intercalary growth makes the leaf bases longer to match the hypopodium's growth, creating a leaf-like sheath with veins around the tuber. In the subsequent growing season, the apical meristem of the dormant bud becomes active, giving rise to an aboveground shoot and a root system that develops from an unusual location. The parent tuberous section of the previous season eventually depletes its reserves and shrinks, likely providing mainly starch reserves to subsequent plants[45].

    • A previous study by Le Roux & Robbertse[21] revealed that Gloriosa superba L. seeds with the sarcotesta intact never germinated and were heavily contaminated by fungi. After the sarcotesta was removed, seeds germinated at an ambient temperature of 23 ± 2 °C, achieving a germination percentage of 21.5%. However, germination was significantly lower at 30 °C and nonexistent at 35 °C. Germination remained irregular across all temperature ranges, with the highest germination rate observed at 23 ± 2 °C after 31 d[21]. These findings suggest that Gloriosa superba L. seeds germinate best at ambient temperatures, and the removal of the seed coat significantly alleviates physical dormancy. However, this intervention does not guarantee the alleviation of morphological seed dormancy, as overall germination rates remain poor regardless of temperature.

      This study examined how different plant growth regulators (GA3, BAP, NAA, and KN) affected the germination of Gloriosa superba L. seeds. The most effective treatments for breaking seed dormancy and commencing germination in Gloriosa superba L. seeds were the application of 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP. Germination started as early as the 3rd day after the seeds were treated with these substances. The treatment exhibited the highest mean seed germination throughout all recorded time intervals (days 3, 5, 9, 13, 21, and 30), resulting in an overall survival percentage of 77.08% by the 30th day. In addition, the application of different concentrations of GA3 in combination with KN and NAA led to survival rates of 72.92% and 68.75%, respectively. The results of this study are the first to show that using the right amounts of GA3 and cytokinin, specifically BAP, together is the best way to break the dormancy of Gloriosa superba L. seeds and help them germinate. This treatment outperforms the effects of GA3 when combined with KN or NAA. Numerous earlier research projects have shown that cytokinins and gibberellins work together to make tomatoes grow faster[47]. Giving Nannochloropsis oceanica cytokinin and GA3 together led to more growth, more lipid production, and more polyunsaturated fatty acids and eicosapentaenoic acid being made. This highlights the potential of this combined effect in various plant species[48]. It has been shown that the interaction between cytokinins and gibberellins has a big impact on cell division and cytodifferentiation. This shows how important this interaction is in many areas of plant biology[49]. Surprisingly, the seedling survival rate obtained during the germination experiment was unexpectedly lower compared to the remarkable survival rate attained during the seed sterilization experiment. However, it should be noted that the seed survival rates in the germination experiment were calculated based on a 30-d duration. This unexpected observation suggests potential factors, such as abiotic stresses, that could have impacted seedling growth beyond the first 30 d. Additionally, the meticulous separation, measurement, and use of different seed batches in this study could explain the variations in seedling survival rates due to disparities in seed viability.

      Seed soaking is a well-established technique for enhancing germination rates by softening the seed coat and removing germination-inhibiting substances[22]. Additionally, pre-treatment with 0.15% HgCl2 for 8 min during sterilization can chemically scarify seeds and induce molecular changes due to the entry of Hg2+ during water imbibition. This chemical scarification helps break seed dormancy and promotes more efficient germination. As noted previously, the use of sterilants not only sterilizes seeds but could also potentially aid germination by neutralizing germination inhibitors[33,43]. The results of this study suggest that an efficient method of seed sterilization can weaken the seed coat and indirectly promote chemical scarification. This is a necessary step for speeding up the process of breaking seed dormancy when combined with a specific formulation of GA3 and other plant growth regulators, thus improving the effectiveness of seed germination (Fig. 6). In a prior study comparing the efficacy of potassium nitrate and gibberellic acid in breaking seed dormancy in both coated and uncoated seeds of Agrimonia eupatoria L., researchers found that the treatments were more effective in promoting seed germination in uncoated seeds compared to coated ones. Furthermore, the study indicated that combining seed coat removal with potassium nitrate application not only increased seed germination rates but also enhanced seedling length[50]. The authors hypothesized that potassium ions from potassium nitrate enhanced cell wall permeability, leading to heightened enzyme activity and cellular metabolism. Their research also suggested that potassium nitrate might help seeds germinate by changing the balance of hormones inside the seed, which could lower inhibitors like abscisic acid and end physiological dormancy[50]. Another prior study highlighted contrasting responses in Vigna radiata (mungbean) when exposed to mercury stress, with seedling growth and root elongation proving more sensitive compared to seed germination[51]. Mercury treatments across all concentrations significantly reduced seed germination, shoot and root length, and seedling dry weight relative to the control, indicating a detrimental impact on mungbean germination and growth[51].

      Recent studies underscore the critical role of aquaporins in seed imbibition and subsequent germination[52]. It has been demonstrated that abiotic stresses, including water deficit, salinity, and heavy metals, modulate the expression of aquaporins across different plant organs[53]. These membrane proteins are particularly abundant in regions of cell division and enlargement, making them critical for facilitating water transport between adjacent cells during seed germination. Seed aquaporin activity closely links to water uptake and the initiation of metabolic processes within the seed, thereby promoting enhanced germination rates[52]. Furthermore, heavy metals such as cadmium (Cd), copper (Cu), and mercury (Hg) can significantly impact the gene expression and function of D-myo-inositol-3-phosphate synthase (MIPS) in plants[53]. MIPS is essential for de novo inositol synthesis and is highly expressed in developing seeds. It plays a critical role in stress responses and regulates the synthesis of phytate, the major phosphorus storage compound in seeds. Moreover, MIPS is integral to complex plant stress response mechanisms and participates in various biochemical and physiological processes, including intracellular signal transduction, membrane construction, protein anchoring, cell wall construction, and auxin storage and trafficking within plant seeds[53]. Arabidopsis possesses three MIPS genes, with MIPS1 being the most extensively characterized. Researchers have shown that a loss-of-function mutant in mips1 causes deformed cotyledon development[54]. Among the three genes, MIPS1 exhibits the highest expression during seed development. Double mutants, mips1 mips2+/− and mips1 mips3, display severe embryogenesis defects, leading to altered cotyledon numbers and deformed shapes. Homozygous triple mutants are embryonically lethal, underscoring the critical role of de novo synthesis of myo-inositol in proper development[54].

      These findings highlight the differential impact of heavy metal ions on pathways associated with seed germination and seedling growth. While germinating seeds are highly sensitive and their germination can be inhibited by heavy metals like mercury, which block aquaporins, these metals also can modulate gene expression and the function of myo-inositol phosphate synthase (MIPS) in various plant species. The potential blocking effect of aquaporins in seeds due to prolonged sterilization in mercuric chloride could have been mitigated or negated by the in vitro growth conditions. The gel-like consistency of the media plays a crucial role in regulating water absorption by seeds by retaining water, maintaining surface moisture, aiding nutrient transport, and ensuring a stable pH. This controlled environment is essential for seed germination and early seedling growth. Furthermore, the density of the gel in plant tissue culture media significantly influences seed germination rates by managing water absorption and moisture levels. In this study, the gel density was optimal, as excessively dense gels can limit oxygen availability and gas exchange, while moderate gel loading enhances germination speed and synchrony. Achieving the optimal gel density was critical for balancing moisture and aeration, thereby ensuring optimal seed growth and germination[55]. Furthermore, depending on the plant species and genotype, this modulation can potentially bypass the aquaporin-blocking effect, resulting in either positive or negative impacts on seed germination and seedling growth[52]. For instance, in Pisum sativum, the reduction of root hydraulic conductivity (Lpr) due to HgCl2 treatment was accompanied by an increase in the expression of plasma membrane intrinsic protein (PIP), suggesting a compensatory mechanism for the blocked aquaporins. Conversely, in Populus deltoides roots subjected to copper stress, genes encoding plasmalemma (PIP) and tonoplast (TIP) aquaporins were downregulated under Cu application[52]. Although the specific effect of HgCl2 treatment on MIPS in Gloriosa superba L. seeds was not evaluated through molecular studies, it is plausible that Hg ions played a critical role in upregulating MIPS pathway-related genes in these seeds. This upregulation could have significantly influenced seed germination, thereby enhancing subsequent seedling growth and development.

    • The present study aimed to evaluate the impact of different combinations of various types and concentrations of plant growth regulator (PGR) treatments on seedling growth parameters. Under carefully controlled in vitro conditions, seedlings grew much faster after being treated with plant growth regulators (PGRs) as compared with the group that did not receive PGR treatment. It was observed that adding 1.5 mg·L−1 BAP and 1.0 mg·L−1 NAA to MS media with 30 g·L−1 sucrose and a 16-h photoperiod enhanced seedling growth after 4 weeks of culture. This treatment significantly improved seedling length, seedling root length, and seedling biomass. This result shows that, when used at the right concentrations, the combined impact of BAP and NAA on the growth of Gloriosa superba L. seedlings is greater than that of 2iP and IAA (Fig. 8af). A previous study on Cymbidium aloifolium L. found that adding BAP and NAA to the MS medium accelerated seedling growth compared to when the medium lacked plant growth regulators[56]. For Scutellaria bornmuelleri, the optimal plant growth regulator combination for growth and development was TDZ and BAP[57]. Conversely, 2iP and IAA were the most effective in achieving direct organogenesis. This highlights the varied impacts that different types and combinations of plant growth regulators can have on the growth and developmental processes of different plant species[57].

    • The experiment on seedling growth showed that the best combination of plant growth regulators (1.5 mg·L−1 BAP and 1.0 mg·L−1 NAA in MS media with 30 g·L−1 sucrose and a 16-h photoperiod) made many seedling traits much better. The only trait that wasn't significantly better was the shoot-to-root ratio, which was the same as the control (Fig. 8c). The seedling roots showed a non-adventitious characteristic, with an average length of 4.08 cm, and had limited growth. This observation fits with how the species usually germinates in the hypogeum, which is to focus on the first growth of the tuberous hypopodium while the roots of the seedling grow. These tubers serve as crucial nutrient stores for the plant, housing inactive buds that subsequently become active, resulting in the emergence of new shoots and root systems. Therefore, following the results of the seedling growth experiment, it was determined in this study to excise the shoots of the fully grown young seedlings. Subsequently, these shoots underwent a rooting treatment to promote further root growth before proceeding with the hardening procedure.

      Several studies have looked at how auxin-group hormones (IAA, IBA, and NAA) affect how roots form and how plants grow in general[58]. This study examined the effect of auxin hormones on root induction and development in microshoots derived from in vitro-grown Gloriosa superba L. seedlings. This study emphasized the efficacy of 1.0 mg·L−1 IBA for rooting. This concentration exhibited the fastest rate of root formation and the shortest time for root initiation, and it performed exceptionally well in terms of average root length and the number of roots per explant. This observation aligns with the findings documented in prior investigations[59,60]. The process of shoot excision and rooting demonstrated exceptional efficacy. Although the average root length yielded comparable outcomes to the seedling development experiment, this method resulted in the growth of robust adventitious roots. Significantly, the quantity of roots produced per shoot was considerably greater in comparison to the initial seedling development experiment (Fig. 10e).

    • Hardening is a crucial stage in the multiplication of tissue culture plants. It entails gradually acclimating plantlets from laboratory settings to soil, which signifies the ultimate achievement of the procedure. The plantlets, which had developed strong shoots and roots, were withdrawn and thoroughly washed to remove any agar residue. They were then transplanted into vessels filled with a sterile mixture of vermiculite and soil in a 1:1 volume ratio. The containers were initially placed in a controlled culture environment for 2 weeks, during which they adapted to their new substrate. Afterward, they spent an extra week in a net house with shade, where they were planted in a mixture of garden soil, sand, and vermiculite at a ratio of 2 parts soil to 1 part sand to 1 part vermiculite (volume/volume). Surprisingly, all plantlets survived without exception during this time of acclimation. However, after being transplanted onto the field around 11 weeks later, the survival rate decreased to 60%. According to Mosoh et al.[29], the main reason for the drop in survival rates is that the plantlets are exposed to a lot more photosynthetically active radiation (PAR) when they move from vessels with garden soil, sand, and vermiculite in the shaded net house to the open field. Moreover, it is possible that the three-week time adjustment was insufficient to provide the required resilience for immediate transplanting into the field. This situation corresponds to the more general occurrence observed when plants produced in a controlled environment are transferred to soil and exposed to conditions outside of the controlled environment. This can lead to reductions in plant survival rates due to changes in the environment that have a negative impact[29]. Notably, there were no noticeable variations in the physical or developmental traits of the surviving plantlets.

    • In this study, sterilizing with 0.15% mercuric chloride (HgCl2) for 8 min removed all contaminants and had an amazing 100% survival rate for the seedlings. This led to the production of seeds that were free of pests and had strong seedling growth. The best treatment used Murashige and Skoog (MS) medium with 1.5 mg·L−1 GA3 and 1.5 mg·L−1 BAP, along with 4% sucrose and a 16-h photoperiod. This resulted in great seed germination (9.25 out of 12 seed explants) and a great overall seedling survival rate of 77.08% after a month. Also, seedlings that were 2 weeks old were grown on MS medium with 1.5 mg·L−1 BAP and 1.0 mg·L−1 NAA, 30 g·L−1 sucrose, and a 16-h photoperiod. After 4 weeks, the seedlings were the longest (5.83 cm), and the roots were the longest (4.08 cm). Moving cut seedling shoots to half-strength MS medium with 1.0 mg·L−1 IBA increased the rate of root formation by 84.37% and helped roots grow even more. Subsequently, these in vitro-grown plantlets were successfully acclimatized and transplanted under field conditions, achieving a commendable 60% survival rate after 11 weeks.

      Effective sterilization of seeds is crucial for removing contaminants and facilitating important processes such as seed coat softening and chemical scarification. Integration with plant growth regulators accelerates the breaking of seed dormancy, thereby enhancing germination rates. Precise seed treatment protocols are essential for advancing seed propagation and conservation practices in plant science. This study provides a robust methodology for the mass in vitro propagation of Gloriosa superba L., alleviating strain on the species' wild populations. Additionally, evaluating supply chains, market dynamics, and prospects is likely to benefit both small-scale stakeholders and the phytopharmaceutical industry.

      In the future, it is crucial to prioritize research that thoroughly investigates the complex mechanistic elements underlying the effectiveness of sterilization in breaking seed dormancy. This research should prioritize elucidating the intricate molecular mechanisms responsible for seed dormancy release and the enhancement of germination facilitated by heavy metal ions, such as Hg2+, and potentially other metals. Understanding these mechanisms is pivotal for advancing our comprehension of how environmental factors impact seed physiology and germination processes, with implications spanning agriculture, conservation, and ecosystem management. By delving into seed biology at a molecular level, such studies will lay the groundwork for developing precise treatment protocols. This, in turn, promises to elevate germination success rates and refine propagation strategies, thereby bolstering efforts in sustainable agriculture and biodiversity conservation.

    • The authors confirm their contribution to the paper as follows: study conceptualization and design: Khandel AK; data collection and curation: Mosoh DA, Khandel AK; analysis and interpretation of results: Mosoh DA; funding Acquisition: Mosoh DA, Khandel AK, Vendrame WA; investigation: Mosoh DA, Khandel AK, Verma SK, Vendrame WA; methodology: Mosoh DA, Khandel AK, Verma SK; project management: Mosoh DA, Khandel AK; resources: Mosoh DA, Khandel AK; software: Mosoh DA; supervision: Mosoh DA, Khandel AK, Verma SK, Vendrame WA; validation: Mosoh DA, Khandel AK, Vendrame WA; visualization: Mosoh DA; original draft manuscript preparation: Mosoh DA; revised – manuscript: Mosoh DA, Vendrame WA. All authors reviewed the results and approved the final version of the manuscript.

    • The data that support the findings of this study are available on request from the corresponding author.

      • The USDA National Institute of Food and Agriculture has provided support for this study, specifically under Hatch project 7001563. In the early stages of the project, Dr. Rohit Sharma from the Centre for Biodiversity Exploration and Conservation (CBEC) provided crucial support, for which we are grateful. His contributions have played a crucial role in determining the course of our research. We express our profound gratitude to Mr. Tetu Acha Samuel (MD, USA) and his colleague, Mr. Adamou Musa (TX, USA), for their prompt assistance with appropriation.

      • The authors declare that they have no conflict of interest.

      • Received 29 May 2024; Accepted 29 July 2024; Published online 12 September 2024

      • Gloriosa superba L. is a critically endangered tropical plant with significant medicinal, cultural, and ornamental value. However, seed dormancy represents a significant barrier to natural regeneration in wild plant populations.

        This study found that sterilizing seeds with 0.15% mercuric chloride (HgCl2) for 8 minutes effectively eliminated contamination, ensuring 100% seedling survival.

        Pre-treatment with HgCl2 weakened the seed coat and induced chemical scarification, enhancing in vitro seed germination. Moreover, the imbibed Hg2+ ions likely induced the upregulation of MIPS genes in the seeds, which subsequently complemented the action of plant growth regulators (PGRs) in enhancing seed germination, seedling growth, and development.

        The combination of GA3 and BAP was most effective for in vitro seed germination. The combination of BAP and NAA was most effective for seedling development and elongation, while IBA was the best for rooting microshoots derived from seedlings.

      • Copyright: © 2024 by the author(s). Published by Maximum Academic Press on behalf of Hainan University. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (12)  Table (4) References (60)
  • About this article
    Cite this article
    Mosoh DA, Khandel AK, Verma SK, Vendrame WA. 2024. Overcoming dual seed dormancy and enhancing in vitro seedling development of Gloriosa superba (L.) with a targeted sterilization approach and plant growth regulator synergy. Tropical Plants 3: e031 doi: 10.48130/tp-0024-0033
    Mosoh DA, Khandel AK, Verma SK, Vendrame WA. 2024. Overcoming dual seed dormancy and enhancing in vitro seedling development of Gloriosa superba (L.) with a targeted sterilization approach and plant growth regulator synergy. Tropical Plants 3: e031 doi: 10.48130/tp-0024-0033

Catalog

  • About this article

/

DownLoad:  Full-Size Img  PowerPoint
Return
Return