Cannabis sativa: From Therapeutic Uses to Micropropagation and Beyond

4.2.1. Direct Organogenesis of Cannabis sativa

Direct organogenesis refers to the process whereby organs are formed directly on the surface of the cultured explant, bypassing the need for a callus phase. Plant regeneration via direct organogenesis involves various steps: initiation of shoot bud; shoot proliferation; shoot elongation and rooting. This type of plant regeneration is preferred over indirect organogenesis as it avoids unwanted somaclonal variation. Direct regeneration makes use of meristematic tissues and various protocols have been established for propagation of C. sativa by use of direct organogenesis via axillary buds, shoot tips and cotyledons as explants [50,51,52] (refer to Table 1).

Several tissue culture protocols established for C. sativa have demonstrated the efficient use of different plant growth regulators (PGRs) (Table 1). Lata et al. [50] studied various effects that different concentrations of CK [kinetin (KN), thidiazuron (TDZ) and benzyladenine (BA)] have on the proliferation of nodal explants using axillary buds. Of these CKs, the highest rate of shoot induction was observed with TDZ application when concentrations of 0.11 mg/L, were used. Even though using cytokinins alone is adequate for shoot multiplication, some studies suggest that addition of low concentrations of auxin may in some cases be beneficial [53]. This being said, Wang et al. [51] examined the effects of BA, TDZ, and KN on bud formation in shoot tip explants with or without naphthaleneacetic acid (NAA) and reported a high plantlet response in medium supplemented with 0.2 mg/L TDZ and 0.1 mg/L NAA. The frequency of plantlet regeneration had a bud multiplication rate of 3.22 per shoot tip. An efficient protocol for the micropropagation of C. sativa using meta-Topolin (mT), a novel aromatic cytokinins has been reported [54,55]. Lata et al. [54] developed a one-step protocol promoting shoot formation and root induction. The study reported a 100% shoot induction at 0.49 mg/L mT concentration. A protocol by Kodym et al. [56] used industry-based fertiliser (Canna Aqua Vega Fertilizer A + B Set, The Netherlands), rockwool blocks and forced ventilation, without the need for PGRs, sugar or vitamins. From this study, the authors reported a 95% shoot induction and rooting rate while the stock culture could be maintained for 6 months. Medium supplementation with TDZ (0.11 mg/L) resulted in a 100% culture response, with an average of 13 shoots per explant [50].

Another protocol includes a patent by Grace et al. [57] that reports a method of significantly producing pathogen-free plants and also pathogen-free clones that comprise the heating of a progenitor plant to alternating temperatures of approximately 100 and 85 °F before surface sterilisation with bleach solution. This patented protocol exploits the fast regeneration capacity of plant meristems where the excising of meristematic tips and their transfer onto MS culture medium for further culturing lead to high induction rates for organogenesis. Although the number of patented protocols is few for Cannabis, another micropropagation system was patented by Hari [58]. This patent provides a regimen for the generation of new Cannabis varieties with modified phytochemical and growth profiles. The innovation described therein subjects plant parts to pectinase digestion (1 mg/mL of pectinase in isotonic buffer, incubated for 3 h) to release plant cells. Following this, the cells are then centrifuged and the cells in the form of a pellet are then cultured on MS medium or Gamborg B5 callus culture media. A second patent by Hari [59] presents a method in which the released plant cells are suspended in a mutagenic solution to obtain mutated Cannabis cells prior to their growth on culture medium. Although these protocols are innovative, they can become labour intensive and costly when enzymatic steps are required as part of the procedures of recovering explants for tissue culture. This also means that highly skilled personnel are needed in order to generate digested cells that are still viable that will continue to grow without abnormalities in culture. Tissue culture steps that are simple may be easier to setup in an industrial platform designed for mass multiplication of plants. One of the limiting factors in the tissue culture of Cannabis variants is associated with use of different strains, making developed protocols more difficult to establish when another cultivar is being micropropagated. This is illustrated by Codesido et al. [60], where organogenesis responses were difficult to predict even when the same medium was being used. Axillary buds are often used in tissue culture due to their highly regenerative nature and survival rates can even be at 100% (Table 1) but the choice of PGR being used must be of high priority. For example, the use of mT proved better for plantlet regeneration compared to the addition of NAA and IBA for direct organogenesis [60].

The explant choice can limit tissue culture growth productivity, and, in some cases, the use of shoot tips and nodal segments yields lowers regeneration frequencies, with explants failing to become highly prolific in their growth patterns [61]. In the study by Galán-Ávila et al. [47], the response of the hypocotyl sections was preferred in comparison to cotyledons when the medium was supplemented with ZEA^RIB at 2 mg/L. The addition of 0.02 mg/L NAA was similar to the medium containing the cytokinin alone as a frequency of plantlet regeneration was recorded at 66.67%. Hyperhydricity can be problematic in tissue culture, leading to plants that are abnormal in their growth and development and such cultures are generally more difficult to acclimatise [62]. The use of vented jars that allow for a better gaseous exchange is one method to minimise the hyperhydration of cultured microshoots to generate acclimated plants rapidly that are healthy. Somaclonal variation is kept to a minimum and even avoided by direct organogenesis protocols [54]. Many of the protocols presented were the first of their kind. Nodal explants using mT, an aromatic natural cytokinin (cytokinin N⁶–(3–hydroxybenzyl) adenine), have been tested for their successful application on in vitro propagation of Cannabis sativa, using a one-step protocol promoting shoot formation and root induction in the same medium [54]. This study also reported that 100% of explants placed on a medium with 0.49 mg/L mT produced shoots. The first report of the use of hypocotyls as explants was reported in the work of Galán-Ávila [47]. Piunno et al. [48] reported the first known shoot regeneration protocol from floral tissues in this species.

meta-Topolins are often regarded as a better alternative to replace BA in culture. meta-Topolins as a supplement in micropropagation of C. sativa should be explored further following the success of the protocol presented by Lata et al. [54]. Galán-Ávila et al. [47] and Piunno et al. [48] both reported new explant types that have not been studied before, hypocotyl and floral tissues, respectively. Because of the novelty of these protocols, these should be further investigated to assess efficiency. These explants could also be tested in protocols that have already proven to have high efficiency. The use of seeds as explants seems to be a popular choice and can help avoid pests but requires constant attention when growing as they can develop into male plants. Seeds may be the cheapest method, but seeds need to be feminised prior to germination to ensure there are no male plants (https://www.royalqueenseeds.com/blog-how-to-make-feminised-cannabis-seeds-like-the-pros-n1117 (accessed on 15 December 2020)). Using clones serves as a rapid alternative and guarantees plants with desired genetics. The plants grown from seeds take longer and may not produce identical clones, which is a disadvantage when the intention is to generate plants that are similar in their clonal fidelity.

Temporary immersion of plant cultures using both semi-solid and liquid cultures has the potential to increase plant yields, allowing for ease of scale-up whilst producing microcultures with minimal symptoms of physiological disorders. Various types of temporary immersion systems are available but thus far the RITA® system has been the one tested for Cannabis using shoot tip culture and nodal explants (Table 1) and compared to a self-designed jar in the study by Kodym and Leeb [56]. Although both temporary immersion bioreactors showed good plantlet regeneration, the ease of handling of the plant material during subcultures was thus preferred by the authors. It is likely that other innovations for use of a variety of different temporary immersion bioreactors will be tested in future studies.

A best-practice regimen may be difficult to obtain for plants that are inherently high-producers of phenolics as wounding that is required for explant sectioning during tissue culture leads to increased production of phenolics as part of the wounding response. The manual handling of the plant material during decontamination steps may also cause physical injury that exacerbates phenolic exudation from the explants. The inclusion of activated charcoal is a common practice as it decreases toxic metabolites, substantially preventing the onset and excessive accumulation of phenolic compounds that may lead to unprecedented explant mortality. For Cannabis, several authors have used charcoal as a preventative measure to control phenolics in culture so as not to compromise the health of microplants. The studies of Piunno et al. [48], Grulichová and Mendel [63], Lata et al. [54], and Lata et al. [50] illustrate the application of charcoal for this purpose but, concomitant to this, charcoal leads to the adsorption of PGRs, changing the intracellular phytohormone balances of plants, encouraging the establishment of root initials and extension of lateral roots [64].

4.2.2. Indirect Organogenesis of Cannabis sativa

Indirect organogenesis involves shoot regeneration from morphogenic callus. Callus-mediated organogenesis depends on various factors including PGRs and explant type. Shoots, roots, and plant formation from callus cultures can be achieved through the manipulation of PGRs in the medium. Young leaves, petioles, internodes and axillary buds were amongst the most commonly used explants for indirect organogenesis in C. sativa (Table 2). Callus tissue are made up of various cell types and the formation of meristemoids relies greatly on the culture medium and also the plant growth regulators applied to this medium. Callus formation is generally induced by increasing cytokinin concentration in the medium and decreasing the concentration of auxins in the medium. Ślusarkiewicz-Jarzina et al. [68] supplemented MS medium with various PGRs to induce callus using a concentration range of 1–4 mg/L kinetin, NAA at 0.5–2 mg/L, 2,4-D (2–4 mg/L) or dicamba (DIC) ranging from 2 to 3 mg/L and petiole explants responded with the highest frequency of callus formation recorded at 82.7% when the growth media had 2–3 mg/L DIC. Although the generation of the callus exhibited high rates, the conversion of this callus to microplant propagules was disappointing, with low organogenesis being problematic. Contrary to this, Wielgus et al. [69] generated callus that could be induced from all types of explants, with cotyledon explants showing the highest callus induction efficiency, and stem explants showing the highest plant regeneration rate. DARIA medium containing NAA, kinetin and BA proved to be an efficient regimen for C. sativa plant regeneration [55] and was used to generate in vitro growth of Cannabis sativa L., resulting in higher callus induction in comparison to prior studies.

To optimise in vitro callus induction and regeneration of Cannabis, Movahedi et al. [70] investigated the efficiency of BA and TDZ included in the medium at 0.1–3 and 0.1–3 mg/L, respectively, on epicotyl and cotyledon explants to generate callus. These two PGRs were used individually or in combination with 0.5 mg/L IBA. This PGR combination generally leads to direct organogenesis but results from this study showed that callus formation was dominant over direct regeneration, contrary to what is usually expected [70]. A rapid protocol for C. sativa plantlet production from young leaf tissue was developed by Lata et al. [71]. Culture initiation took place on MS medium + 0.09–0.35 mg/L IAA, 0.1–0.41 mg/L IBA, 0.09–0.37 mg/L NAA, 0.11–0.44 mg/L 2,4-D with 0.22 mg/L TDZ, while shoots were induced on MS medium + 0.11–2.25 mg/L BAP 0.12–2.15 mg/L KN, and 0.11–2.2 mg/L TDZ. Callus formation was achieved in medium supplemented with 0.09 mg/L NAA and 0.22 mg/L TDZ, whereas the highest shoot induction and proliferation were obtained from 0.11 mg/L TDZ. It was, however, shown that the use of TDZ in plant cell culture might not be the best option due to high levels of DNA methylation in callus cultures that potentially lead to somaclonal variation [72]. Such variations are defined by higher levels of polymorphism, affecting clonal fidelity. Callus formation can be very heterogeneous and can vary tremendously between different types of explants, so many studies test various explants to test their success. Raharjo et al. [73], similarly to Lata et al. [71], investigated the response of leaves as explants and compared leaf material to the use of flowers and seedlings. Interestingly, in that study, leaves were productive in callus formation and flowers were highly effective, giving the most callus in culture. The generation of callus from seedlings was regarded as advantageous and valuable for Cannabis regeneration studies.

Page et al. [74] emphasised the influence of different basal media in their ability to encourage the production of callus, hyperhydricity and low rates of microplant production and the importance of exploring other basal media in addition to MS. In that study, of the five genotypes tested, the MS medium led to higher incidences of callus production when combined with TDZ at 0.11 mg/L but a greater leaf canopy was established when this DKW medium was used instead of MS [75]. The in vitro response of the four different genotypes of medicinal Cannabis that were similar may thus indicate the potential to obtain a more standardised protocol that can be used to reliably propagate a wide range of Cannabis strains in vitro. Many plant tissue culture protocols for Cannabis rely on the use of leaf material or seedlings for plantlet regeneration, even though flowers as an explant source have been shown to be beneficial in other plant species that exhibit recalcitrance to shoot organogenesis. The use of a floral reversion strategy that converts florets to vegetative tissues, under a 12 h photoperiod, led to significantly better shoot development rates in comparison to the use of apical and axillary nodal explants [76]. This method of plantlet production in vitro for Cannabis offers an innovative and reproducible technique that could easily be adopted for the commercial production of different Cannabis cultivars especially when mT at 0.24 mg/L is used as the PGR in the medium [76].

The literature on micropropagation of Cannabis, whether it be for the botanical drug markets or other industrial sectors, is a growing scientific concern and future endeavours are likely to result in more innovative approaches to support traditional micropropagation schemes that are currently existing [77]. Although much research has been targeting solutions in finding appropriate growth media and PGR combinations that can produce normal and healthy plantlets of C. sativa cultivars, a shift in the research foci of biotechnologists is becoming more obvious. For example, there is an increasing body of literature where physical parameters are being evaluated for their effects on micropropagation and this is a change in thinking, as many previous studies that populate the literature were more focused on finding plant growth regulator combinations that could elicit more prolific shoot regeneration in vitro.

Nowadays, the influence of physical conditions that may control organogenetic responses in tissue cultured plants of Cannabis, especially those that are associated with light conditions, is becoming more apparent. Ventilation of the culture vessels for micropropagation in Cannabis and other plant genera is thought to be important in producing true-to-type plantlets that are can be readily acclimatised [67,78]. The benefits of investigating physical factors are many. Some of these include Cannabis plantlets that are better able to maintain photosynthesis, grow in microenvironments with reduced humidity, and perform gaseous exchange more efficiently, thereby closely resembling plants that are grown ex vitro [67]. To investigate the influence of physical factors, Murphy and Adelberg [67] tested the hedging technique, which involves the removal of the apical meristem, over several subcultures using C. sativa ‘US Nursery Cherry 1’, and in this study, the medium was used without the inclusion of PGRs to prevent genetic drift over several continuous culture passages. In non-vented culture jars, higher light intensities were important to maintain a vigorously growing and regenerating culture stock but ventilation in vitro, allowing for better gaseous exchange, assisted with the production of healthier plantlets which could easily be acclimated that also had lower plantlet mortalities ex vitro. These authors iterated the strong apical dominance that is exhibited by Cannabis in vitro, which ultimately leads to poor multiplication and shoot branching rates. Therefore, decapitating the apical meristematic tissues thus offers a solution which can promote shoot branch formation and multiplication for Cannabis and when this is combined with high light intensities (at 120 μmol m⁻² s⁻¹ PPFD), multiplication rates can improve significantly. Cutting off the apical meristem is easily doable during subculture and should thus be taken into great consideration for a species that is associated with poor multiple shoot cluster formation in a plant tissue culture setting.

As reproducibility of published techniques for micropropagation of other genotypes, apart from the MX Cannabis variety, is a growing concern, Monthony et al. [45] tested 10 drug-type variants (genotypes GRC, RTG, U22, U31, U37, U38, U42, U61, U82 and U91) in an attempt to replicate the study by Lata et al. [54]. A genotype-specific response for callus production was evident, with many of the tested genotypes showing browning of the callus due to the accumulation of phenolics. Although callus could be generated for the test genotypes using MS supplemented with a combination of 0.22 mg/L TDZ and 0.09 mg/L NAA, the conversion of the callus to regenerated plantlets was deemed difficult as low plantlet regeneration rates were observed across all the tested Cannabis varieties in that particular study regimen. This is a major problem for the establishment of reproducible protocols that can be commercially used as new Cannabis-bred lines become more available in years to come.

4.2.3. In Vitro and Ex Vitro Rooting of Cannabis sativa

At present, several different PGR regimes are available for use as part of rooting steps that are implemented for Cannabis cultivars. In the rooting stage, explants are induced to form roots typically by the application of an auxin. Indole-3-butyric acid (IBA) and indole-3-acetic acid (IAA) are amongst the most used PGRs for root induction. High rooting frequencies were obtained by combining a synthetic auxin such as NAA with a natural auxin such as IAA (Table 1 and Table 2) [55]. meta-Topolins have been reported to stimulate in vitro rooting in some plant species without the need for auxin application—in turn, these conditions improve survival rates in acclimatised plants [79]. Some of the methods that are commonly used include a study performed by Wrobel et al. [61] in which they compared the effects of MS medium supplemented with or without different concentrations (0.25–0.75 mg/L) of either IBA or IAA. However, this study showed no significant difference between these two auxins in terms of the rooting rates of microplants. Contrary to the use of plant growth regulators, shoot tip cuttings that are placed directly into rockwool and into a growth chamber promote rooting and rooting rates of 97.5% of the in vitro shoot tip cuttings allow for successfully acclimatised plants within 3 weeks [56,61]. Lata et al. [54] showed that the best concentration for rooting was 0.05 mg/L mT and that mT concentrations higher than 0.97 mg/L were inhibitory to rooting. Grulichová and Mendel [63] used the same rooting medium as used by Lata et al. [50], whereby full-strength MS medium was supplemented with IBA and activated charcoal. These authors indicated that the highest number of roots were present on plants cultured on media without any PGRs present or on a medium containing 0.5 mg/L mT. Lata et al. [50] recorded a 94% response of cultures rooted in a medium supplemented with 0.51 mg/L IBA, with an average of 4.8 roots per explant and confirmed that activated charcoal is effective in root induction.

Plantlets grown in medium supplemented with mT successfully rooted well in vitro [54]. Ślusarkiewicz-Jarzina et al. [68] transferred plantlets to a rooting medium comprising of MS basal medium supplemented with 1.0 mg/L IAA + 1.0 mg/L NAA, and 69.9% of plantlets formed roots. To induce rooting, Parsons et al. [65] tested NAA and IBA in four concentrations (0.1, 0.2, 0.5 and 1 mg/L) using regenerated shoots. Of the media explored for rhizogenesis, 0.1 mg/L IBA exhibited the highest rooting rate. Lata et al. [71] similarly showed that shoots rooted best in ½ MS medium with 0.51 mg/L IBA. The presence of IBA resulted in a significantly higher rooting percentage (80–96%) than PGRs IAA or NAA.

The rooting of plants in vitro does assist with acclimatisation, often shortening this particular ex vitro growth phase and this is beneficial when plants are targeted for large-scale agricultural production so as to minimise the acclimation steps in the greenhouse prior to field transplantation.

The quality of the end products and, in commercial production, economic viability is determined by the successful ex vitro acclimatisation of micropropagated plants [80]. Greenhouse and field conditions have significantly lower relative humidity and higher light levels compared to in vitro conditions, that prove to be stressful to micropropagated plants. The benefits of a micropropagation system can, however, only be realised by successfully transferring plantlets from tissue-culture vessels to the ex vitro conditions [81].

Conventional ex vitro acclimatisation and rooting encompass the steady weaning of plantlets from culture conditions towards ambient light and humidity levels. Direct and indirect regenerated C. sativa plantlets are subjected to ex vitro rooting and acclimatisation (Table 3). The combinations that are currently used include black peat, granulated peat moss and perlite [47], sterilised soil [61], coco natural growth medium and sterile potting mix-fertilome [54], autoclaved organic manure, clay soil and sand [52] and vermiculite and plant ash [51]. Unlike most soils, cocopeat/coco natural growth medium is unfertilised. This then means that when using this type of media for acclimatisation of plants, an external nutrient solution must be applied, and the pH also needs to be controlled and remain close to neutral. Often the growth of Cannabis variants is discussed via online blogs, for example, refer to Mr. Grow It (https://www.mrgrowit.com) [82].

Cocopeat is the preferred medium amongst growers due its many benefits such as its airy and eco-friendly structure that allows roots to thrive in an oxygen-rich environment. Soil, however, retains water better than cocopeat and often already has naturally occurring nutrients that are then assimilated from soil to plant using nutrient acquisition strategies that are readily employed by plants. In such cases, this then minimises time spent watering and preparing nutrient solutions by the grower. The downside, however, is that the roots do not receive as much air as they would in cocopeat, and good air flow to the roots is needed for the stronger and more rapid growth of the Cannabis plants. This then makes cocopeat the better choice in terms of increasing yield for the purpose of ex vitro rooting.

In most studies, growth room temperature ranges from 22 to 30 °C with 60% relative humidity [47,50,54]. Wrobel et al. [61] reported a 95% survival rate in the growing chamber and a 90% survival rate in field conditions. Plants propagated with mT rooted better when transferred to soil than the shoots produced with TDZ, and plantlets showed a 100% survival rate in acclimatised plants [54]. Lata et al. [50] observed new growth after 2 weeks of ex vitro rooting. Plants reached 14–16 cm in height within 6 weeks of transfer and plants showed normal development and no gross morphological variation. Plantlet survival for 3 months after field transfer was also reported by Wang et al. [51] and the rates were at high levels at 99%.