Introduction

Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize is a fast-growing, temperate-adapted member of the Poaceae (Hodkinson et al. 2002). The potential of M. × giganteus as a sustainable, non-food, lignocellulosic feedstock for bioethanol (Lewandowski 1998), and value-added product development (Engbers and Deen 2013) has been previously demonstrated. Its perennial nature allows for high yields of aboveground biomass for 20–25 yr depending on location and agronomic practices (Xue et al. 2015). After 3 yr of initial establishment, annual biomass yields can range between 31 and 61 t ha−1, which is significantly greater than other biomass crops, such as maize (Zea mays L.) or switchgrass (Panicum virgatum L.) (Heaton et al. 2008). Net profit for yields can amount to as much as $2900 (USD) ha−1 annually 10 yr after initial establishment (Heaton et al. 2004).

Despite these advantages, adoption of M. × giganteus for commercial production is limited, in large part due to establishment costs. M. × giganteus is a seedless triploid (3x = 57) that can be propagated using direct rhizome planting (DRP) or micropropagation with estimated costs of approximately $3835.39 (USD) and $7181.99 (USD) ha−1, respectively (Xue et al. 2015). While micropropagation using current techniques is more expensive than DRP, plantlets derived from somatic embryos (embryoid plants—EP) exhibited greater shoot lengths and tiller number per plant (51 and 19.1 for DRP; 57 and 47.1 for EP, respectively), in the first year of field establishment in Germany (Lewandowski 1998). Morphological characteristics, such as the number of branches over 15-cm long and number of bamboo-like shoots, were greater in EP (7.0 and 11.6, respectively) than DRP (3.9 and 5.5, respectively; Lewandowski 1998). In addition to these characteristics, EP produced twice the shoot biomass and the EP rhizomes that were 135% heavier than DRP in the first year of planting (Lewandowski 1998). Furthermore, in vitro technologies can facilitate genetic improvement of this sterile species through mutagenesis or genome editing (Mehrotra et al. 2007), production of certified disease-free germplasm for national and international exchange (Taşkın et al. 2013), rapid multiplication of “elite” cultivars that exhibit genetic integrity (Rambaud et al. 2013), and aid in the study of plant processes without interference from unwanted artifacts (Vreugdenhil et al. 1998). As such, in vitro technologies offer many advantages for M. × giganteus, but additional research is needed to improve the technology.

Indirect regeneration of various species of Miscanthus has been successfully developed and has mainly focused on somatic embryogenesis from immature inflorescences (Lewandowski 1997; Petersen 1997; Kim et al. 2012; Gubišová et al. 2013; Perera et al. 2015; Ślusarkiewicz-Jarzina et al. 2017). Somatic embryo development in M. × giganteus is typically induced with 2,4-dichlorophenoxyacetic acid (2,4-D; Kim et al. 2012) and low levels of 6-benzylaminopurine (BAP; Lewandowski 1998). A major challenge in M. × giganteus micropropagation is the loss of competence during prolonged callus culture. Kim et al. (2010) attempted to improve regeneration of M. × giganteus plantlets from immature inflorescence cultures. Six weeks after initiation of callus, the majority were classified as yellow/white compact with shoots (41 ± 4%), followed by K2 (22 ± 2.1%), and K3 (37 ± 3.7%) callus tissue. The regeneration frequency of the callus was greatest for shoot-forming and embryogenic-like callus after 1 (0.93) and 4 mo (0.9) of culture on solid callus medium, respectively. However, regeneration was not achieved from callus maintained for more than 4 mo (Kim et al. 2010).

Like other grass species, such as Sorghum bicolor (L.) Moench (sorghum), the regenerative capacity of callus maintained on callus maintenance medium over long periods of time may be hindered by an accumulation or change in phenolic acid content (Liu et al. 2015), which leads to recalcitrance and/or toxic effects to plant tissues (Zaid 1987), modification of endogenous phytohormone levels (Březinová et al. 1996), control of dormancy in somatic embryos, and influence over organogenesis (Cvikrová et al. 1998). Changes in phenolic content may be due to an upregulation of the phenylpropanoid biosynthetic pathway or degradation of lignin macrostructures in response to wounding (Zawadzki and Ragauskas 1999). In M. × giganteus stems, Le Ngoc Huyen et al. (2010) demonstrated that cell walls were largely composed of a 3:1 ratio of ester-linked p-coumaric and ferulic acids. Regeneration of plantlets from M. × giganteus callus culture could be hindered due to formation of premature lignin from these soluble phenolic compounds in callus cells and/or somatic embryos.

Phenol and lignin biosynthesis has been shown to be significantly reduced in in vitro American elm (Ulmus americana L.) suspension cultures, sugar maple (Acer saccharum Marshall) callus, and Artemisia annua L. callus cultures by the addition of 2-aminoindan-2-phosphonic acid (AIP) to the culture media (Jones et al. 2012; Jones and Saxena 2013). 2-Aminoindan-2-phosphonic acid competitively inhibits phenylalanine ammonia lyase (PAL), the enzyme responsible for the first committed step of the phenylpropanoid biosynthetic pathway that converts phenylalanine to ammonia and trans-cinnamic acid (MacDonald and D’Cunha 2007). In A. annua callus cultures, it was found through both visual inspection and measurement of total phenolic content that tissue browning decreased in a dose-dependent manner up to 10-μM AIP, and that 100-μM AIP significantly reduced total tissue phenolics (Jones and Saxena 2013). In addition, total soluble phenolic content has been demonstrated to be correlated with the amount of lignin found in somatic embryos and callus in vitro (Cvikrová et al. 2003), and embryo growth and development were negatively correlated with total phenolic content (Malá et al. 2000). Similar results have been observed in a number of species in vitro, including maize, sessile oak (Quercus petraea Liebl.), and alfalfa (Medicago sativa L.) (Lozovaya et al. 1996; Cvikrová et al. 1998, 1999, 2003; Hrubcová et al. 2000).

The objective of the current study was to optimize the regeneration capacity of M. × giganteus callus after prolonged storage on semi-solid medium. It was hypothesized that the application of AIP would reduce the total soluble phenolic content in callus and subsequently improve regeneration potential and timeframe. The findings of this research may help to improve germplasm conservation, maintain disease-free stocks, and create a foundation for future in vitro breeding technologies.

Materials and Methods

Plant material

Immature inflorescences (5.0–15.0 cm in length), were collected from field-grown M. × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘Illinois’ (‘M161’) at All Weather Farming Incorporated (Port Ryerse, Ontario, Canada) (42° 47′ N 80° 12′ W) on September 15, 2015. Through visual inspection using a modified Biologische Bundesantalt, Bundessortenamt, and CHemische Industrie (BBCH) scale, the majority of plants grown at this location were in principal growth stage 4: booting; specifically, varying between stages 43 to 47 (Tejera and Heaton 2017). Stems were cut beneath the most distal (youngest) node with the sheath intact. Segments were wrapped in moist paper towels and kept in darkness at 4°C for 5 d. Thereafter, all but two to three of the surrounding leaves were removed from the immature inflorescences. Immature inflorescences were rinsed under running tap water for 5 min, soaked in 70% (v/v) ethanol for 1 min, and rinsed with sterile, deionized water three times. Immature inflorescences were then immersed in 20% (v/v) commercial bleach (12.5% sodium hypochlorite; Clorox®; The Clorox Company, Oakland, CA), with approximately 0.1% (v/v) Tween® 20 (Thermo Fisher Scientific® Company, Ottawa, Canada) for 20 min with gentle shaking, followed by rinsing five times with sterile, deionized water. Spikelets (each approximately 5.0 mm in length) were aseptically dissected from the rachis, then inoculated individually for callus induction.

Callus induction and multiplication

Callus induction and maintenance medium consisted of Murashige and Skoog (MS; Murashige and Skoog 1962) basal salts and vitamins (PhytoTechnology Laboratories®, Shawnee Mission, KS), supplemented with 30-g L−1 sucrose, 11.3-μM 2,4-D, 2.2 μM BAP, 1000-mg L−1 L-proline, 300-mg L−1 casein hydrolysate, and 7-g L−1 agar (Kim et al. 2012). All plant growth regulators and L-proline were purchased from Sigma-Aldrich® (St. Louis, MO); casein hydrosylate from PhytoTechology Laboratories®; and Select Agar™, powder from Thermo Fisher Scientific®. Callus induction medium was adjusted to pH 5.7 using 1-N NaOH and 1-N HCl (Sigma-Aldrich®) before autoclaving at 121°C and 144.8 kPa for 20 min. Ten explants were placed on 20 mL of medium in 100 × 15-mm Petri dishes (Thermo Fisher Scientific®). The calluses were incubated in the dark in a growth room at 24 ± 2°C for the duration of the experiment. Callus tissue was subcultured and multiplied monthly (approximately every 28 d) for 6 mo. In the fifth month of culture, calluses measuring 3.0 to 8.0 mm in diameter were inoculated on one of five callus maintenance media. Media were composed as described previously and supplemented with either 0-, 1-, 10-, 100-, or 1000-μM AIP. In the sixth month of culture, half of the calluses from each of these treatments were placed on callus maintenance media modified with 9.0-μM 2,4-D and 0-, 1-, 10-, 100-, or 1000-μM AIP. At each subculture time point, visibly necrotic and/or atrophied tissue was gently removed from callus.

Callus morphology assessment

At each subculture (beginning after the initial 6 mo of culture initiation and multiplication), calluses were visually assessed for morphology using the following rating system: (1) “compact white” callus (K2), characterized by a smooth, white surface (Petersen 1997), generally accompanied by the appearance of “embryogenic-like” structures; (2) “yellow/green” callus (K1), previously described as nodular and moderately soft; (3) “friable” callus (K3), identified by a soft, watery appearance (Kim et al. 2010) usually lacking embryogenic capacity (Morrish et al. 1987); and (4) “browning” callus. Representations of different callus morphologies may be found in Fig. 1a–d.

Figure 1
figure 1

Representative Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ callus morphologies. Calluses were assessed using a dissecting microscope and were putatively classified as: (a) compact yellow/green (K1); (b) somatic embryo forming and friable (K2 and K3, respectively, occurring in the same sample); (c) root forming; and (d) shoot forming. Leaf primordia (e) and differentiated shoot-like structures (f) formed on calluses. The root-caps of regenerated roots were identifiable by the production of red/purple pigmentation and lack of root hairs (g), and anthocyanin spots were noticeable on the remainder of the callus (h). Scale bars represent 1 mm.

Full size image

Development of differentiated and embryo-like structures

In addition to callus morphology, the following characteristics were recorded: the development of “shoot-like” and “root-like” structures (herein referred to as “shoots” and “roots,” respectively) and the formation of “embryo-like” structures (herein referred to as “somatic embryos”) (representations may be found in Fig. 1e–h). The frequency of each callus type and accompanied structures was calculated as callus type or structure (%) = (number of each callus type or structure ÷ total number of calluses plate−1) × 100 (Kim et al. 2010).

Shoot, root, and somatic embryo counts were conducted using a dissecting microscope. Analyses of differentiated structures were conducted between 9 and 13 mo of culture, while somatic embryo counts began at 11 mo of culture. Individual leaves on the callus constituted distinct shoots. Since somatic embryos typically developed in clusters and were quantified without destruction of the callus, conservative counts were conducted from the visible surface of the callus. For the duration of the AIP treatment period, six calluses were placed on each medium with five replications each. Care was taken to group calluses of similar morphologies on the same plate.

Soluble phenolic content

The soluble phenolic content of calluses from each treatment was measured after 2 mo on AIP-supplemented media, with sampling occurring monthly for 5 mo. At each subculture, one callus (with necrotic and/or atrophied tissue removed) was removed from each treatment replicate (five replicates per treatment) and pooled in a 15-mL centrifuge tube (Thermo Fisher Scientific®). The sample g fresh weight (FW) from each treatment was recorded before flash-freezing in liquid nitrogen, followed by lyophilisation using a FreeZone™ 4.5 freeze dryer (Labconco®, Kansas City, MO). After lyophilization, the sample g dry weight (DW) was recorded, total moisture content (MC) (%) was calculated [((FW-DW) ÷ FW) × 100], and samples were finely ground. Extraction solvent was added to each tube (80% methanol, v/v), with a 1:10 tissue to solvent ratio (w/v). The tubes were vortexed and placed in a sonicating water bath (Branson 3510, Danbury, CT) for 30 min. The tubes were then removed and centrifuged for 10 min at 21.1 ×g using a Fisherbrand™ AccuSpin™ Micro 17 (Thermo Fisher Scientific®). The supernatant from each sample was transferred into a new microcentrifuge tube.

The soluble phenolic content was assessed using a modified Folin and Ciocalteu (F-C) colorimetric assay. A standard curve was constructed using gallic acid (Sigma-Aldrich®) to obtain gallic acid equivalent (GAE) values at concentrations of 31.25, 62.5, 125, 250, 500, and 1000 μg mL−1 (Folin and Ciocalteu 1927). In brief, 10-μL aliquots of sample extracts, standards, or sample blanks were added to each well of a 96-well flat bottom microplate (Corning, Corning, NY). A volume of 100 μL of 1:10 water: F-C phenol reagent (MP Biomedicals, Santa Ana, CA) was added to each well, and the plate was incubated for 5 min before adding 80 μL of aqueous 0.25 M Na2CO3 (Sigma-Aldrich®). The plate was then incubated in the dark for 1 h before absorbance values at 725 nm were measured with a Synergy™ H1 microplate reader (Biotek, Winooski, VT). All sample and standard readings were corrected with blanks, and all samples, standards, and blanks were replicated in triplicate.

Regeneration and in vitro plantlet development

At 6 mo of culture, excess shoot-forming callus from each 11.3-μM 2,4-D treatment measuring 3.0–8.0 mm in diameter were placed on regeneration medium described by Kim et al. (2012). Regeneration medium consisted of MS basal salts and vitamins supplemented with 30-g L−1 sucrose, 5-mg L−1 BAP, 1-mg L−1 2,4-D, 1000-mg L−1 L-proline, 300-mg L−1 casein hydrolysate, and 7-g L−1 Select Agar™, powder. Regeneration medium was adjusted to pH 5.8 using 1-N NaOH and 1-N HCl before autoclaving at 121°C at 144.8 kPa for 20 min. Six to seven calluses were placed on each plate and incubated in the dark for 1 mo. After regeneration, calluses with shoots were transferred to either semi-solid root induction [MS basal salts with 30-g L−1 sucrose, 0.1% (w/v) activated charcoal (Sigma-Aldrich®), and 7-g L−1 Select Agar™, powder, pH 5.8], or liquid tillering medium [MS basal salts with vitamins, 30-g L−1 sucrose, 5-mg L−1 BAP, 0.1-mg L−1 indole-3-buytric acid (IBA; Sigma-Aldrich®), and 0.45-mg L−1 indole-3-acetic acid (IAA; Sigma-Aldrich®), pH 5.7] for plantlet development (Kim et al. 2012). The cultures were incubated in a growth room at a temperature of 24 ± 2°C, under a 16-h photoperiod with light intensity of 40 μmol m2 s−1 provided by cool-white fluorescent bulbs (Philips® Canada, Scarborough, Canada). All chemicals listed above that are not followed by a supplier were purchased from the same companies as those described in “Callus induction and multiplication.”

Experimental design and statistical analysis

Data obtained from non-destructive visual assessments [“callus morphology frequencies” (7 mo), “number of structures,” “phenolic content,” and “total MC” (5 mo), and “number of embryo-like structures” (3 mo)] were subject to variance analysis conducted using a mixed-model repeated measures analysis of variance (ANOVA), using PROC GLIMMEX in SAS® 9.4 (SAS Institute Inc., Cary, NC). The experiment was constructed in a factorial design and arranged in a completely randomized design (CRD). Each treatment was replicated over five plates, each containing six samples. A logit link function with beta distribution was used for assessing callus morphology frequencies, and an identity link with lognormal distribution was used to analyze the number of differentiated structures and somatic embryos. Correlation coefficients (R2) regarding the occurrence of differentiated structures and somatic embryos on different callus types were calculated. For phenolic content and total MC, each analyzed treatment was composed of five individual calluses from five replicate plates. A type I error rate of α = 0.05 was assigned for all analyses.

Results

Callus morphology frequencies

The percentage of calluses producing shoots and roots were significantly affected by interactions between culture time and AIP concentration. For shoot-forming calluses, declines were observed as culture time progressed in all individual treatments (Fig. 2). Apart from media lacking AIP in the first month of treatment (66.7 ± 6.811%), the greatest frequency of calluses developing shoots was observed on media supplemented with 10-μM AIP (peaking in the third culture month at 58.3%). Regarding the proportion of calluses producing shoots compared to root and somatic embryos, 100-μM AIP supplemented with 11.3-μM 2,4-D exhibited the highest value by the third culture month (73.3%).

Figure 2
figure 2

Proportion of Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ calluses exhibiting shoots, roots-, and somatic embryos over time. Data are presented by 0 (a and b), 1 (c and d), 10 (e and f), 100 (g and h), and 1000 (i and j) μM 2-aminoindan-2-phosphonic acid (AIP) treatments grouped by 9.0 (a, c, e, g, and i) and 11.3 (b, d, f, h, and j) μM 2,4-dichlorophenoxyacetic acid (2,4-D). Each bar represents the mean of 30 samples.

Full size image

The highest frequency of rooting calluses was observed on medium supplemented with 10-μM AIP (45.4 ± 4.663%), followed by 1- and 100-μM treatments (24.5 ± 3.705 and 25.2 ± 3.467%, respectively). Compared to calluses producing other distinctive structures, 1000-μM AIP supplemented with 11.3-μM 2,4-D accounted for 60 to 63.3% root-forming calluses by the sixth and seventh culture mo.

Three-way interactions between AIP concentration, 2,4-D concentration, and culture time were observed with somatic embryo-forming calluses. When the proportion of somatic embryo calluses were compared to shoot- and root-forming calluses, higher values were generally observed in AIP treatments supplemented with 9.0 than 11.3-μM 2,4-D.

The frequencies of K1, K2, and K3 calluses, along with those which browned and could not be properly classified, are depicted in Fig. 3. Differentiated structures and somatic embryos were able to develop on every callus type; however, the occurrence of shoots on K2 and K3 calluses was significantly correlated (Table 1). Additionally, the development of somatic embryos was positively correlated with K2 calluses, while negatively correlated with K1 and K3 types.

Figure 3
figure 3

Proportion of Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ compact yellow/green (K1), somatic embryo forming (K2), friable (K3), and browning callus over time. Data are presented by 0 (a and b), 1 (c and d), 10 (e and f), 100 (g and h), and 1000 (i and j) μM 2-aminoindan-2-phosphonic acid (AIP) treatments grouped by 9.0 (a, c, e, g, and i) and 11.3 (b, d, f, h, and j) μM 2,4-dichlorophenoxyacetic acid (2,4-D). Each bar represents the mean of 30 samples.

Full size image
Table 1 Correlations regarding the occurrence of structures (shoots, roots, and somatic embryos) on the following Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ callus types: compact yellow/green (K1), somatic embryo forming (K2), friable (K3), and browning callus. Sample correlations (R2) followed by an asterisk (*) are significant at α = 0.05
Full size table

Differentiated structure and somatic embryo counts

AIP concentration significantly influenced the average number of shoots per callus, with 10 μM AIP exhibiting the highest counts per callus (4.2 ± 0.5464), followed by 100- and 1000-μM treatments (3.8 and 3.2, respectively) (refer to Fig. 4 for mean structure counts averaged over time and Fig. 5 for means at each mo).

Figure 4
figure 4

Average count of (a) shoots, (b) roots, and (c) somatic embryos developing on Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ calluses cultured on different levels of 2-aminoindan-2-phosphonic acid (AIP) and averaged over time. Each bar represents the mean of 300 (a and b) and 180 (c) samples ± S.E.

Full size image
Figure 5
figure 5

Average count of (a) shoots, (b) roots, and (c) somatic embryos per Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ callus at specific culture times. The legends associated with each graph represent culture time (approximately 1 mo). Each bar represents the mean of 60 samples ± S. E (pooled).

Full size image

The average number of roots demonstrated significant interactions between culture time and AIP concentration. The highest values for the average number of roots were observed in the fifth culture month in treatments 1- and 100-μM AIP and the fourth month in 1000-μM AIP (17.3 and 13.2 ± 2.444, and 18.8 ± 3.5655, respectively). Increasing trends for root development were observed in 1-, 100-, and 1000-μM AIP treatments as time progressed throughout 5 mo of culture, while 0- and 10-μM treatments remained relatively unchanged during this time span.

Although the average number of somatic embryos ranged from 14.9 (1000-μM AIP) to 33.7 (100-μM AIP), no observable trends were detected with these levels when analyzed over the culture period.

Soluble phenolic content

Total soluble phenolic content (μg g−1 DW) was negatively associated with increasing AIP concentrations, with the highest content being observed at 1 μM (2242.3 ± 149.03-μg g−1 DW), and the lowest at 1000-μM AIP (1569.7-μg g−1 DW) (Fig. 6).

Figure 6
figure 6

Average gallic acid equivalent (GAE) (μg g−1 dry weight (DW)) values of Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ calluses cultured on media supplemented with various 2-aminoindan-2-phosphonic acid (AIP) levels. Each bar represents the mean of 50 calluses over two 2,4-dichlorophenoxyacetic acid (2,4-D) levels (10 callus samples over five replications) ± S.E. Bars labeled with the same letter are not significantly different (p < 0.05) according to Tukey’s honest significant difference (HSD) test.

Full size image

Total MC (%) was significantly affected by both 2,4-D and AIP concentrations; however, there were no interactions between these two factors (p = 0.1893). Total MC was generally greater in 11.3-μM 2,4-D and 1000 μM-AIP media compared to the remaining treatments (Table 2).

Table 2 Total moisture content (MC) (%) of Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ calluses used for soluble phenolic content measurements. Analyzes were conducted separately for 2-aminoindan-2-phosphonic acid (AIP) and 2,4-dichlorophenoxyacetic acid (2,4-D). Each mean represents 50 and 100 calluses (10 and 25 callus samples over five replications, respectively). Means followed by the same letter are not significantly different (p < 0.05) according to Tukey’s honest significant difference (HSD) test
Full size table

In vitro plantlet development

After calluses were maintained on callus maintenance media for 6 mo, a portion of the shoot-forming samples from each AIP treatment (cultured on media supplemented with 11.3-μM 2,4-D) were transferred to regeneration medium. After 1 mo, regenerants were transferred to semi-solid rooting- or liquid tillering-media. Shoot-forming calluses from all AIP treatments were able to develop healthy roots and/or shoots in vitro (representations may be observed in Fig. 7). Shoot multiplication rates ranged from six to 18 per explant (data not shown).

Figure 7
figure 7

Regenerated Miscanthus × giganteus J.M.Greef, Deuter ex Hodk., Renvoize ‘M161’ calluses in rooting medium (a), liquid tillering medium (b), and after removal from tillering medium after approximately 28 d of incubation (c).

Full size image

Discussion

Regeneration of M. × giganteus through somatic embryogenesis and/or shoot organogenesis from immature inflorescences has been developed previously (Lewandowski 1997; Głowacka et al. 2010a; Kim et al. 2010; Kim et al. 2012; Gubišová et al. 2013). While this approach offers opportunities for genetic improvement and large-scale plant propagation of this sterile triploid, several problems have been encountered that limit its application. Specifically, callus has been reported to lose its regenerative capacity within 4 mo of culture (Kim et al. 2010), which presents issues for applying this technology. In the current study, AIP—a potent inhibitor of the phenylpropanoid pathway—was found to increase the percentage of calluses exhibiting regeneration of shoots over relatively-extended culture periods.

The capacity for shoot and somatic embryo development from M. × giganteus callus (the majority of these structures occurring on K2 callus) was achieved after prolonged culture (7 mo on AIP-supplemented media; 12 mo including the callus induction phase) on semi-solid callus maintenance media for both 2,4-D concentrations tested. The results presented here are an improvement from the findings reported by Kim et al. (2010), which showed complete loss of shoot-regenerative and embryogenic-development capacity after 4 mo of culture on semi-solid media. Though the current study recognized that the control treatment (0-μM AIP) was able to produce shoot-forming calluses exceeding 4 mo of culture, this could be attributed to a number of factors, including potential differences in genotype, immature inflorescence explant developmental stage used in callus induction (Głowacka et al. 2010b), and/or more care taken to remove necrotic/atrophied tissue at each subculture. Nonetheless, this system will allow for a relatively long term, low-maintenance, effective regeneration system for M. × giganteus ‘M161’, better known as the “Illinois” variety (Withers 2015).

The frequency of different callus types, differentiated structures, and somatic embryos was highly variable among and within treatments, and the proportions of each of these changed noticeably over the duration of the experiment. This phenomena have previously been observed in other grasses, such as Saccharum spp. (sugarcane) (Fitch and Moore 1990). Additionally, the presence of browning callus could have been classified as any of the typical callus morphologies reported in this study; however, tissue necrosis due to any number of stressors made its original classification indiscernible.

Somatic embryos were identified by a smooth, rounded architecture, which appeared either white or green in color. These structures varied in shape, with clusters of embryos represented by a conglomerate of spherical (globular)- and torpedo-shaped structures. While histological analysis to verify their bipolar nature was not conducted, previous studies using a similar system demonstrated that these structures exhibited a bipolar configuration with a discernable epidermis, which is characteristic of somatic embryos (Ślusarkiewicz-Jarzina et al. 2017).

The occurrence of differentiated structures on the various callus types reported in this study is generally in agreement with what has been documented in the literature. Shoots (determined by the presence of green buds or primordia and light green, translucent leaves) formed primarily on K2 (Gairi and Rashid 2004) calluses. Leaves could be ascertained by the presence of trichomes observable on all parts of the tissue, including the leaf tip. Previously reported by Owaga (2015), who worked on the regeneration of Switchgrass (P. virgatum L.) cultures, two callus morphologies were described: type I, which was characterized as a compact, nodular, white to light-yellow callus and type II, being friable, semi-soft, and highly regenerable (Vasil and Vasil 1994). Based on these definitions, the K3 calluses described in this study fit the characteristics of type II callus, which may explain the significant correlation of shoot occurrence on K3 (R2 = 0.56, p < 0.0001).

Calluses that developed roots were generally nodular and semi-soft in appearance (K1) and occurred alongside anthocyanin spots similar to what has been previously reported in Miscanthus (Petersen 1997). These structures were glabrous and displayed a red/purple pigmentation in the area of the calyptra. In previous grass culture studies, K1 calluses are generally root-forming (Morrish et al. 1987); however, the current study did not observe a strong correlation (R2 = 0.22, p = 0.1164) between the occurrence of roots and K1 calluses.

The modified F-C assay (Singleton and Rossi 1965) used in this study is a rapid, non-specific colorimetric technique, which functions to quantify the amount of readily oxidized phenolic substances (Singleton et al. 1999) within biological material. In this study, soluble phenolic content was significantly affected by AIP levels, with the lowest and highest values detected in tissues cultured on media supplemented with 1000- and 1-μM AIP, respectively. Previous authors found that supplementing 10-μM AIP in the medium of alfalfa (M. sativa L.) cultures increased mitotic activity and prompted cellular division (Hrubcová et al. 2000). This has also been demonstrated in sessile oak cultures, in which increased total phenolic acid content positively correlated with non-converting embryos, and decreased phenolic content through application of AIP yielded embryos that more readily regenerated into plantlets (Cvikrová et al. 1998). In the current study, AIP failed to increase the capacity for regeneration in M. × giganteus over time compared to the control; however, the percentage of calluses exhibiting regenerated shoots increased significantly in 10- and 100-μM AIP. Regeneration of calluses at the highest concentration (1000-μM AIP) was lower than samples cultured on these two AIP treatments, and this was accompanied by a greater total MC (87.3%). Similar side effects caused by drastic inhibition of the phenylpropanoid biosynthetic pathway have been documented previously and include increased incidence of hyperhydricity caused by reduced lignification and reduced lignin production for cell wall formation and development of shoots (Cvikrová et al. 2003). This could be of particular significance for grass species, as phenylpropanoids are major components of their cell walls and play important roles in cell growth/development and intercellular adhesion (Jones and Saxena 2013).

Another deleterious effect of high AIP levels in other systems is the hindrance of root formation and growth, as demonstrated in Secale cereale L. (Reuber et al. 1993). In contrast, average root development peaked in calluses incubated on 1000-μM AIP in the fourth culture month (18.8). The frequency of calluses that exhibited root regeneration was more strongly represented in cultures maintained on media with higher auxin levels, which could be explained by the adventitious rooting promoting effects of 2,4-D (Viana and Mantell 1998). As with shoot regeneration, roots were induced for the extent of the culture period, although investigation of root culture with roots derived from callus tissue requires further examination in M. × giganteus.

Conclusions

The results presented here demonstrate that the regeneration capacity of M. × giganteus can be maintained for relatively long-term culture on semi-solid callus maintenance media, and that the percentage of calluses demonstrating regeneration can be enhanced with the addition of 10- and 100-μM AIP. 2-Aminoindan-2-phosphonic acid functions to inhibit the phenylpropanoid biosynthetic pathway and thus can mediate phytohormone levels, cell wall development, and auxin metabolism (Hrubcová et al. 2000). Although soluble phenolic content decreased in a dose-dependent manner with AIP, a more targeted approach at specific phenolic content changes, including cell wall-bound phenols, and to account for potential interfering compounds could contribute to a better understanding of the regeneration capacity of this species. The findings of this study will help to develop options for alternative propagation methods through somatic embryogenesis and synthetic seeds and help to develop a foundation for in vitro breeding strategies.