Skip to main content

Metabolic engineering of energycane to hyperaccumulate lipids in vegetative biomass



The metabolic engineering of high-biomass crops for lipid production in their vegetative biomass has recently been proposed as a strategy to elevate energy density and lipid yields for biodiesel production. Energycane and sugarcane are highly polyploid, interspecific hybrids between Saccharum officinarum and Saccharum spontaneum that differ in the amount of ancestral contribution to their genomes. This results in greater biomass yield and persistence in energycane, which makes it the preferred target crop for biofuel production.


Here, we report on the hyperaccumulation of triacylglycerol (TAG) in energycane following the overexpression of the lipogenic factors Diacylglycerol acyltransferase1-2 (DGAT1-2) and Oleosin1 (OLE1) in combination with RNAi suppression of SUGAR-DEPENDENT1 (SDP1) and Trigalactosyl diacylglycerol1 (TGD1). TAG accumulated up to 1.52% of leaf dry weight (DW,) a rate that was 30-fold that of non-modified energycane, in addition to almost doubling the total fatty acid content in leaves to 4.42% of its DW. Pearson’s correlation analysis showed that the accumulation of TAG had the highest correlation with the expression level of ZmDGAT1-2, followed by the level of RNAi suppression for SDP1.


This is the first report on the metabolic engineering of energycane and demonstrates that this resilient, high-biomass crop is an excellent target for the further optimization of the production of lipids from vegetative tissues.


Advanced biofuels are expected to supply 70% of aviation fuel and 50% of the fuel used in freight transport by 2060 [1]. Biofuel enhances the reliability, security, and affordability of the energy supply and reduces carbon emissions to combat global warming [2]. Production of biodiesel or cellulosic ethanol from renewable and perennial feedstocks is expected to result in significant environmental benefit [3]. These fuels are derived from sugars or lipids that are produced in feedstocks via photosynthesis [4,5,6]. Triacylglycerol (TAG) represents the major lipid component of plant seeds, which provides a highly dense source of energy for the germination and establishment of seedlings [7]. Metabolic engineering of high-biomass crops for the hyperaccumulation of lipids in their vegetative biomass has been proposed as strategy to surpass the oil yields of traditional oilseed crops [8,9,10].

Under typical growth conditions, plant leaves can synthesize TAG but do not hyperaccumulate it. For example, sugarcane leaves have a TAG content of less than 0.05% of their leaf DW [10]. The biosynthetic pathway for TAG production is highly conserved across species, and the genes encoding the major catalytic steps involved in fatty acid (FA), glycerolipid, TAG production, and hydrolysis have been identified. WRINKLED1 (WRI1) is a transcription factor and a member of the APETALA2 (AP2)/ethylene-responsive element-binding protein (EREBP) subfamily. It is a positive activator of FA biosynthesis. Overexpression of WRI1 has been shown to increase TAG accumulation in Arabidopsis leaves 2.8-fold [11]. Suppression of trigalactosyldiacylglycerol 1 (TGD1) also results in elevation of the extraplastidial FA pool available for TAG assembly and may provide an alternative to the ectopic overexpression of WRI1 [12]. TGD1 is a putative component of a lipid transporter that transfers lipids from the endoplasmic reticulum (ER) to chloroplasts [13]. The Arabidopsis TGD1 mutant has increased TAG accumulation in leaves [14]. Diacylglycerol acyltransferase1-2 (DGAT1-2) is a rate-limiting enzyme for the conversion of diacylglycerol into TAG [15]. Oleosin1 (OLE1) is a structural protein that protects lipid droplets from coalescence and reduces lipid turnover [16, 17]. A synthetic OLE1 (CysOLE1) with six engineered cysteine residues improves FA contents in vegetative tissues of Arabidopsis [18]. After TAG is synthesized in the ER, SUGAR-DEPENDENT1 (SDP1), a specific TAG lipase, can catalyze its hydrolysis [19]. The suppression of SDP1 has been shown to increase TAG accumulation in vegetative tissues [20, 21]. Similarly, the mutation of a subunit of the peroxisomal fatty acid ABC transporter (PXA1), which contributes to lipid transport across the peroxisomal membrane for β-oxidation, results in an increase in TAG accumulation in expanding leaves [22].

A “push–pull–protect” strategy has been proposed to increase plant lipid content in vegetative tissue [23]. In this strategy, the genes involved in lipid synthesis (push) and TAG assembly (pull) are overexpressed, while lipid turnover is suppressed (protect) through multi-gene engineering. Indeed, TAG accumulation has been found to increase in vegetative tissues of both model plants such as Arabidopsis thaliana, Brachypodium distachyon, Nicotiana benthamiana, and Nicotiana tabacum [23,24,25,26,27] and high biomass crops including sugarcane [10, 28], maize [29], sorghum [30], and perennial ryegrass [31], through the overexpression of WRI1, DGAT1-2, and OLE1 and/or the suppression of SDP1 or PXA1. For example, in one study, the constitutive coexpression of WRI1, DGAT1-2, and CysOLE1 and simultaneous RNAi-mediated cosuppression of ADP-glucose pyrophosphorylase (AGPase) and a subunit of peroxisomal ABC transporter1 (PXA1) elevated TAG accumulation in leaves of transgenic sugarcane to 1.90% of dry weight (DW), compared to 0.02% of DW in non-modified sugarcane [10].

Energycane, like sugarcane, is an interspecific hybrid in the genus Saccharum. By contrast to sugarcane, energycane has a high proportion of the ancestral species Saccharum spontaneum in its genome, which contributes to higher tiller number, greater biomass yield, more fiber content, and better persistence, in addition to reduced stem diameter and sugar content [32]. Energycane is an ideal feedstock for the production of lipids in vegetative tissues, due to its superior biomass production, tolerance to pests and diseases, persistence on marginal land, and elevated cold tolerance [33,34,35,36]. However, it is among the most recalcitrant species in regard to tissue culture and genetic transformation; to date, there have been few reports of successful energycane transformation. Two such reports provided detailed biolistic transformation protocols using neomycin phosphotransferase II (nptII) [37] or bar as a selectable marker gene [38]. A third report described the use of transgenic energycane as a platform for producing a recombinant therapeutic protein via overexpression of a cDNA encoding snowdrop lectin [39]. Here, we report what is to the best of our knowledge the first metabolic engineering of energycane. Following multi-gene engineering for the hyperaccumulation of TAG, the levels of transgene expression/target gene suppression and the TAG and total FA accumulation in leaves of transgenic energycane were analyzed.


PCR analysis of transgenic energycane

PCR primer pairs were designed to amplify both the 5′ and 3′ ends of each of the two cotransformed, unlinked recombinant DNA constructs (Fig. 1; Additional file 1: Table S1) containing the overexpression cassettes of ZmDGAT1–2 and SiCysOLE1 and/or the RNAi suppression cassettes of SDP1 and TGD1, in addition to the nptII selectable marker gene (Fig. 1A, B). Biolistic gene transfer and the regeneration of transgenic plants (Fig. 2) resulted in 31 transgenic lines from 13 shots, as confirmed by NPTII immuno-chromatography and nptII PCR (Fig. 2; Additional file 1: Table S2). Of the 31 transgenic lines, eight (P5, P8, P16, P19, P24, P26, P27, and P28) were confirmed by PCR to contain both of the cotransformed linearized, unlinked recombinant DNA constructs carrying the overexpression cassettes of ZmDGAT1-2 and SiCysOLE1, as well as the RNAi suppression cassettes of SDP1 and TGD1, in addition to nptII (Table 1 and Additional file 1: Table S2 and Fig. S1); these were selected for further quantitative real-time PCR (qRT-PCR) and TAG analyses.

Fig. 1
figure 1

Multi-gene expression and RNAi suppression constructs that were cotransformed into energycane. A The overexpression cassette of ZmDGAT1-2 and SiCysOLE1. B RNAi hairpin cassette of TGD1 and SDP1. Neomycin phosphotransferase II (NPTII) is used as the selectable marker. ZmDGAT1-2: Diacylglycerol acyltransferase1-2 from Zea mays. SiCysOLE1: Cysteine oleosin1 from Sesamum indicum. TGD1: Trigalactosyl diacylglycerol1 from Saccharum spp. hybrid. SDP1: Sugar-dependent1 from Saccharum spp. hybrid. pZmUbi and pSbUbi: Ubiquitin promoters from Zea mays and Sorghum bicolor, respectively. tNos: Nos terminator from Agrobacterium tumefaciens. tSbHSP: The heat shock protein 18 terminator from Sorghum bicolor. p35S and t35S: 35S promoter and terminator, respectively. Half arrows indicate primers used to detect transgenes

Fig. 2
figure 2

Generation of transgenic energycane. A Leaf whorl cross-sections were used as explants. B Callus induced from leaf whorl cross-section. C Calli regenerating on medium with geneticin for selection. D Regenerated plants before transfer to soil. E NPTII immuno-chromatography of crude protein extracts from transgenic energycane lines (P5, P8, and P16) and non-transgenic energycane (WT), arrow NPTII test line, Bar = 1 cm. F Transgenic energycane lines (P5, P8, and P16) next to non-transgenic energycane (WT)

Table 1 Summary of TAG content and expression/suppression of transgenes in leaves of transgenic energycane

Of the 31 transgenic lines, eight were only PCR-positive for the nptII selectable marker and RNAi suppression cassettes. Of the 31 transgenic lines, 15 were PCR-positive only for one of the two PCR reactions that were carried out for each of the two cotransferred recombinant DNA constructs, indicating fragmented inserts. Lines P6, P7, and P20, which were PCR-positive for ZmDGAT1-2; SiCysOLE1, and line P23, which was PCR-positive for ZmDGAT1-2, were also included in the qRT-PCR and TAG analyses (Additional file 1: Table S2). The non-transgenic energycane CP 82-1592 (wild type, WT) was used as negative control. Sugarcane line 1565, described by Parajuli et al. [28], which was grown under the same conditions, was also included in RT-PCR and TAG analyses.

TAG and total FA accumulation in the leaves of transgenic energycane and sugarcane

The transgenic lines had a TAG content ranging from 0.11 to 1.52% of leaf DW, which was 2- to 30-fold of that of WT (Table 1). Sugarcane line 1565, described by Parajuli et al. [28], was also grown under the same conditions and displayed TAG contents of 4.87% of leaf DW (Additional file 1: Table S3). The energycane lines P5, P8, and P16, which had the highest TAG contents at 0.65%, 0.90%, and 1.52% of leaf DW (Fig. 3), respectively, were selected to analyze total FA content as well as TAG and total FA composition. P5, P8, and P16 had a total FA content of 3.56%, 4.74%, and 4.96% of leaf DW, respectively, the latter being almost double that in WT (2.70% of DW) (Fig. 3). The total FA content was highly positively correlated with the TAG content (0.93) in Pearson’s correlation analysis (Additional file 1: Table S4).

Fig. 3
figure 3

TAG and total FA content of transgenic energycane lines (P5, P8, and P16). Values with different letters are significantly different at p ≤ 0.05 according to a one-way ANOVA test and the Duncan’s Multiple Range Test (MRT), administered post hoc. P5, P8, and P16 are lines with the highest TAG content compared to other transgenic lines. WT Non-transgenic energy cane, TAG Triacylglycerol, TFA Total fatty acid, DW dry weight

TAG and total FA composition in leaves of transgenic energycane

The composition analysis of TAG FA revealed that the unsaturated FA, oleic acid (C18:1Δ9), increased from 0.00% in WT to 5.68–10.86% leaf TAG FA in transgenic lines P5, P8, and P16. The content of the saturated FAs, palmitic acid (16:0), and stearic acid (18:0) was reduced in the transgenic lines to 12.97–46.52% of those of WT (Fig. 4A). The amount of linoleic acid (LA, 18:2Δ9,12) was significantly increased in transgenic lines 2.88- to 4.68-fold, at the expense of α-linoleic acid (ALA, 18:3Δ9,12,15) (Fig. 4A). Similarly, in the total FA composition analysis, the content of C18:1Δ9 and 18:2Δ9,12 of the three lines was significantly elevated 1.17- to 2.80-fold compared to WT, while the content of 16:0, 18:0, and 18:3Δ9,12,15 was reduced to 74.98–88.40% of the WT levels (Fig. 4B). The levels of other FAs in all three lines were increased 1.02- to 3.12-fold compared to those of WT in both the TAG and total FA composition analyses (Fig. 4A, B).

Fig. 4
figure 4

TAG composition and total FA composition in leaves of transgenic energycane lines with higher TAG content. A Different TAG compositions in leaves of transgenic lines and non-transgenic energycane (WT). B Different total fatty acid compositions in leaves of transgenic lines and WT. Values with different letters are significantly different at p ≤ 0.05 according to a one-way ANOVA test and Duncan’s Multiple Range Test (MRT), administered post hoc. P5, P8, and P16 are lines with higher TAG content than other transgenic lines. TAG Triacylglycerol, FA fatty acid

Overexpression/suppression levels of transgenes in transgenic energycane and sugarcane

qRT-PCR analysis was conducted to examine the levels of transgene expression and target gene suppression in the split leaves used for the TAG and FA analyses. ZmDGAT1-2 was overexpressed, with a relative expression level ranging from 0.06 to 1.63, in all transgenic lines, except for line P24, which did not express ZmDGAT1-2 (Table 1). SiCysOLE1 was expressed at less than 0.1 relative to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) in transgenic lines, except for line P5, which had an expression level of 0.26 relative to GAPDH (Table 1). TGD1 RNAi resulted in a transcript suppression of up to 18% of WT levels (line P28; Table 1). The suppression of TGD1 was identified in both lines with high TAG contents ranging between 0.65 and 1.52% TAG of DW (P5, P8, and P16) and lines with low TAG content ranging between 0.11 and 0.19% TAG of DW (P24, P25, P26, P27, and P28) (Table 1). Interestingly, most lines with ZmDGAT1-2 and/or TGD1 suppression showed a 5–74% elevated SDP1 expression relative to WT levels despite the cotransformation with an SDP1 RNAi construct. However, the lines that had the highest TAG accumulation (P8, P16, and P5) displayed modest SDP1 suppression, with 95%, 81%, and 70% of WT levels, respectively (Table 1). The transgenic lines that had the highest TAG content (P5, P16, and P8) displayed the highest relative expression level of ZmDGAT1-2 at 0.55, 1.44, and 1.63, respectively. By contrast, lines with low TAG content ranging between 0.11 and 0.19% TAG of DW, such as P19, P24, P25, P26, P27, and P28, displayed a relative ZmDGAT1-2 expression level of less than 0.16 (Table 1). By contrast to energycane lines P5, P8, and P16, sugarcane line 1565 displayed WRI1 expression (0.05 relative to GAPDH) and stronger suppression of SDP1 (30% relative to WT levels), while ZmDGAT1-2 (0.19 relative to GAPDH) and SiCysOLE1 expression (0.02 relative to GAPDH Additional file 1: Table S3) were lower than in the highest-expressing energycane lines.

Correlation between the levels of transgene expression/target gene suppression and TAG accumulation in transgenic energycane

Pearson’s correlation was evaluated for TAG accumulation and the levels of expression of lipid genes in leaves of transgenic lines. As shown in Table 2, TAG accumulation was highly positively correlated with the expression level of ZmDGAT1-2, with a correlation coefficient of 0.86 while it was negatively correlated with the expression level of SDP1 with a correlation coefficient of − 0.52. However, TAG content was not significantly correlated with the expression of SiCysOLE1 or TGD1 in transgenic energycane (Table 2).

Table 2 Correlation of (trans)gene expression with TAG content in transgenic energycane


Energycane produces among the highest amounts of biomass of any crop, which, in combination with its resilience and ease of biocontainment, makes it a prime feedstock candidate for fueling the emerging bioeconomy [33,34,35,36]. However, its recalcitrance in regard to tissue culture and genetic transformation has so far prevented complex metabolic engineering and molecular pharming approaches. To the best of our knowledge, only one earlier report presents energycane as a production platform for recombinant protein [39].

Here we report the first complex metabolic engineering of energycane, which resulted in a hyperaccumulation of lipids in vegetative biomass. Using a multi-gene approach with constitutive overexpression of transgenes ZmDGAT1-2 and SiCysOLE1 and the RNAi suppression of TGD1 and SDP1, transgenic energycane accumulated TAG at 1.52% of leaf DW in an average of three biological replicates. This TAG accumulation exceeds that detected in non-transgenic energycane (WT) by 30-fold (Table 1). The level of TAG hyperaccumulation also resulted in almost twice the total fatty acid content relative to WT (Fig. 3). Similarly, TAG accumulation has been increased through metabolic engineering of lipid biosynthesis genes in vegetative tissues of model plants and crops [10, 23,24,25,26,27,28,29,30,31]. In perennial ryegrass (Lolium perenne), the accumulation of TAG in leaves was increased 14-fold relative to WT to 2.5% of leaf DW when AtDGAT1 and SiCysOLE1 were coexpressed [31]. Interestingly, TAG accumulation was closely correlated to SiCysOLE1 expression and not to AtDGAT1 expression in ryegrass, suggesting that TAG degradation may be higher in ryegrass than in energycane. By contrast, in this work, TAG was most highly correlated with ZmDGAT1 expression and was not correlated with SiCysOLE1 expression. Differences in the source of the DGAT1 gene, codon optimization, and promoter choice, which affected the expression of the individual genes, may also have contributed to these contrasting findings.

ZmDGAT1-2 was also found to be positively correlated with TAG accumulation in transgenic sugarcane lines coexpressing SbWRI1, ZmDGAT1-2, and SiCysOLE1 and suppressing AGPase and PXA1 [10]. While ZmDAGT1 was highly expressed in energycane, SiCysOLE1 expression was very modest in the majority of lines, and the optimization of its expression has the potential to boost TAG accumulation. The SDP1 suppression level was also highly correlated with TAG accumulation in energycane. This finding is in agreement with earlier studies in Arabidopsis, where a mutant background in SDP1 severely decreased FA turnover, leading to more than a twofold increase in leaf TAG accumulation when stacked with WRI1 and DGAT1 overexpression [21] or a 15-fold increase in leaf TAG when stacked with TGD1 suppression [20]. However, TGD1 suppression to 18% of WT levels in energycane was associated with only modest TAG accumulation (0.19% of leaf DW) and therefore did not correlate with TAG accumulation (Tables 1 and 2). Stronger levels of target gene suppression may be facilitated by the gene editing approaches that were recently established for sugarcane [40,41,42,43], in which the highly polyploid nature of sugarcane and energycane offers the opportunity to create a range of knockout levels and phenotypes.

Gene expression and TAG accumulation were also compared directly between the transgenic energycane lines generated in this work and the transgenic sugarcane line 1565, which was reported previously [28]. Sugarcane line 1565 harbored the same gene expression/RNAi suppression cassettes as energycane in addition to a constitutive expression cassette of WRI1. The expression of WRI1 and stronger suppression of SDP1 in sugarcane line 1565 contributed to a 3.2-fold higher TAG accumulation than in energycane line P16 (Table 1 and Additional file 1: Table S3). A synergism between WRI1 and DGAT1 coexpression for TAG accumulation has been reported in tobacco [25]. Similarly, in sorghum, a TAG content of 3–8.4% of leaf DW has been reported for WRI1 coexpressed with DGAT and OLE [30]. These data support the conclusion that transcription factor WRI1 is a critical factor for the hyperaccumulation of TAG. However, the constitutive expression of multiple lipogenic factors can create a certain amount of toxicity to the cell, limiting tissue culture regeneration or compromising plant development and biomass accumulation [27].

The energycane lines P5, P8, and P16, which had TAG accumulations of 0.65%, 0.9%, and 1.52% of leaf DW and lacked WRI1 expression, did not display obvious growth retardation when compared to WT (Fig. 2F; Table 1), while sugarcane line 1565 with expression of WRI1 and TAG accumulation of 4.87% of leaf DW produced only 46% of the biomass of WT [28]. In addition, the tissue culture recalcitrance of energycane may create a bottleneck that leads to the selection of gene expression combinations with lower TAG accumulation levels. Future approaches should include strategies that avoid TAG accumulation in tissue culture or early plant development using stem-specific, inducible, or developmentally regulated promoters. Candidates for stem-specific promoters have been described [44, 45]. The transgenic energycane lines P5, P8, and P16 displayed a substantial increase in the accumulation of unsaturated fatty acids at the expense of saturated fatty acids (Fig. 4A, B). This indicates that the expression of ZmDGAT1-2 preferentially catalyzes the esterification of unsaturated fatty acids to diacylglycerol. This finding is consistent with earlier reports on sugarcane [10, 28] and Arabidopsis [20, 24].

Collectively, the optimization of expression cassettes using developmentally regulated or stem-specific regulatory elements, codon optimization, and stacking of additional lipid biosynthesis-related genes such as DOF4 [46] and a combination with CRISPR/Cas-mediated knockout of genes contributing to lipid hydrolysis could further elevate the accumulation of lipids in energycane.


Energycane is a prime feedstock for the generation of renewable energy and bioproducts owing to its unsurpassed biomass yield and resilience under abiotic and biotic stress. In this work, a multigene expression/suppression strategy produced hyperaccumulation of TAG and total FA in leaves at levels exceeding non-modified energycane by 30- and almost twofold, respectively. Moreover, ZmDGAT1 expression and SDP1 suppression had the highest correlation with TAG accumulation. These results establish energycane as a promising production platform for lipids from vegetative biomass.


Construction of multigene expression vectors

Multigene expression and RNAi vectors were assembled using a conventional restriction enzyme digest of vector components and ligation, as described by Parajuli et al. [28]. In the multigene expression construct, Zea mays ubiquitin promoter (pZmUbi) and the terminator (tNos) of the nopaline synthase gene from Agrobacterium tumefaciens were used in the overexpression cassette of the DGAT1-2 gene from Zea mays (ZmDGAT1-2) (Fig. 1A); the ubiquitin promoter (pSbUbi) and the heat shock protein 18.2 terminator (tSbHSP) from Sorghum bicolor were used to drive the expression of CysOLE1 from Sesamum indicum (SiCysOLE1) (Fig. 1A). For the RNAi suppression of TGD1, an RNAi hairpin was custom synthesized (Genscript, Piscataway NJ) consisting of sense (244 bp) and anti-sense (244 bp) of TGD1 of Saccharum spp. hybrid separated by Paspalum notatum’s 4CL intron (94 bp). Similarly, the RNAi hairpin of SDP1 was custom synthesized (Genscript, Piscataway, NJ, USA) as the sense (278 bp) and anti-sense (278 bp) of SDP1 of Saccharum spp. hybrid separated with a 4CL intron (94 bp) from P. notatum. The hairpins of TGD1 and SDP1 were sub-cloned under transcriptional control of CaMV 35S promoter and terminator (Fig. 1B); the selectable marker gene nptII was placed under transcriptional control of the Zea mays ubiquitin promoter (pZmUbi) and the CaMV 35S terminator (t35S) (Fig. 1B).

Tissue culture and genetic transformation of energycane

After six alternative genotypes were screened, energycane genotype CP 82–1592 [47] was selected for genetic transformation due to its efficiency in embryonic callus induction and plant regeneration. Immature leaf whorl cross-sections (Fig. 2A) were used as explants in the tissue culture, as described by Fouad et al. [37] to induce calli (Fig. 2B); linearized expression cassettes of transgenes were transformed into embryonic calli using the biolistic particle-delivery system; and putative transgenic plants were regenerated after selection with geneticin (Fig. 2C, D). The media used in callus induction, direct embryogenesis, transformation, selection, and shoot and root regeneration were prepared as described by Fouad et al. [37].

For the bombardment, the plasmid was prepared from an overnight culture of 5 mL E. coli strain TOP10 at 37 °C. The backbone of the plasmid was removed using restriction enzyme digestion with AscI. The linearized fragment was gel extracted and purified as described by Fouad et al. [37]. A total of 13 shots of linearized fragment DNA were delivered in a 1:2 molar ratio (for nptII + tgdRNAi + sdpRNAi:DGAT + Ole) to callus using the Biolistic PDS-1000/He particle-delivery system (BioRad, Hercules, CA, USA) as described by Fouad et al. [37]. Rooted plantlets were transferred to a potting mix (Jolly Gardener C/G) after any media residues were washed from the roots and the plants had been covered for 5 days with a Magenta box to provide an environment with high humidity. In the growth chamber, the temperature was controlled at 25–28 °C during the day and 22–24 °C during the night. The photoperiod was set to 16 h light/8 h dark cycles with a light intensity of approximately 400 μmol m−2 s−1.

NPTII immuno-chromatography assay

Leaf tissue from putative transgenic plants was ground in extraction buffer provided with the NPTII ImmunoStrip kit (Agdia Inc., Elkhart, IN, USA) and centrifuged at room temperature for 5 min at 16,100×g; the supernatant was transferred to a new microfuge tube, where it was absorbed by an ImmunoStrip. The presence of NptII was indicated by the development of two purple lines on the immuno-chromatography strip (control and test line), while non-transgenic control plants developed only one purple line (control line) (Fig. 2E).

PCR analysis of genomic DNA extracts

Genomic DNA was extracted from young leaves of regenerated plants using the cetyl trimethyl ammonium bromide method [48]. Then, 100 ng genomic DNA was used as the template in a 20 µL PCR reaction. PCR amplification was conducted using Hot Start Taq DNA Polymerase (NEB; Ipswich, MA, USA) under the following conditions: 95 °C for 30 s, followed by 32 cycles of 95 °C for 30 s; 51.8 °C, 56.5 °C, 57.7 °C, or 58.2 °C for 30 s for TGD1, nptII, CysOle1, or ZmDGAT1-2, respectively; 68 °C for 1 min; and a final elongation at 68 °C for 5 min. The primers for each target gene are listed in Additional file 1: Table S1.

Greenhouse propagation of transgenic, lipid-accumulating energycane

Transgenic energycane (Fig. 2F) was grown in a greenhouse and propagated by nodal stem cuttings to obtain biological replicates and three plants per transgenic line, and non-transgenic plants were each planted in a pot with a 15 cm diameter containing potting mix (Jolly Gardener C/G). Plants were grown under a drip fertigation system. In the greenhouse, the temperature was controlled by evaporation cooling to 25–29 °C during the day and 20–24 °C during the night, using the natural photoperiod with a maximum daily light intensity of approximately 500–1000 μmol m−2 s−1. The plants were irrigated and fertilized via drip fertigation. To compare the lipid gene expression and TAG accumulation between transgenic energycane and sugarcane, non-transgenic sugarcane CP 88–1762 and transgenic sugarcane line 1565, as described earlier by Parajuli et al. [28], were also grown in the same greenhouse in three biological replicates with similar lipid gene expression/suppression cassettes.

Sampling of leaves for qRT-PCR, TAG, and FA analysis

The leaves of the transgenic lines and WT were numbered according to the system proposed by Kuijper et al. [49]. The top visible dewlap leaf blade of the plants that had been growing in the greenhouse for 1 month was used for these experiments. To ensure spatiotemporal correspondence between lipid gene transcripts and TAG and FA accumulation, the dewlap leaf was further divided into two halves at the midrib: one half was used for TAG and FA analyses, while the other half was used for qRT-PCR analysis. One sample was taken from each of the three plants, with a total of three biological replicates from each transgenic line. For the TAG and FA analyses, almost 100 mg fresh leaf tissue was needed for each sample. The samples were immediately freeze-dried in a lyophilizer (Labconco, Kansas, MO, USA) for 72 h. Samples containing lyophilized leaf tissue were delivered to Brookhaven National laboratory on dry ice and were immediately used for the TAG and FA analyses. For the qRT-PCR analysis, 0.1–0.2 g fresh leaf tissue was collected for each sample. The samples were flash-frozen in liquid nitrogen and were subsequently stored at − 80 °C before total RNA extraction.

qRT-PCR analysis of lipid gene expression

Total RNA from leaf samples was extracted using TRIzol reagent (Ambion, Life Technologies, Thermo Fisher Scientific, Waltham, MA, USA); 1.0 µg total RNA from each sample was used for the cDNA synthesis using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster, CA, USA). The gene-specific primers described by Parajuli et al. [28] and shown in Additional file 1: Table S1 were used to evaluate the expression levels of the ZmDGAT1-2, SiCysOLE1, TGD1, and SDP1 genes. The glyceraldehyde 3-phosphate dehydrogenase (GAPDH) gene was used as a housekeeping gene for the normalization of transcripts [50]. qRT-PCR was conducted in a CFX Real-Time PCR detection system (Bio-Rad, Hercules, CA, USA) with SsoAdvanced Universal SYBR green supermix (Bio-Rad, Hercules, CA, USA) according to the conditions provided by Parajuli et al. [28]. The relative transcription levels of ZmDGAT1-2 and SiCysOLE1 were calculated using the 2{Ct (GAPDH)–Ct (transgene)} method. The relative suppression levels of TGD1 and SDP1 were calculated using the 2−ΔΔCt method [51].

Analyses of TAG and total FA composition

TAG and FA analyses were carried out as described by Parajuli et al. [28]. In brief, 700 μL extraction solution in which methanol, chloroform, and formic acid were mixed at a ratio of 2:1:0.1 by volume was added to 10.0 mg lyophilized leaf tissue. After 3 h mixing on a vortex mixer, a hexane:diethyl ether:acetic acid solution (70:30:1 by volume) was added, and the total lipid extracts were divided for thin layer chromatography. Incubated in 1.0 mL boron trichloride-methanol at 80–85 °C for 40 min, TAG fractions were scraped from the plate under UV light and transmethylated into FA methyl esters (FAMEs). In the total FA analysis, after incubation in 1.0 mL boron trichloride-methanol, total lipid extracts were directly transmethylated into FAMES. FAMES were dissolved in 100.0 μL hexane and quantified via gas chromatography-mass spectrometry; 5.0 µg C17:0 was used as an internal standard.

Statistical analysis

The data for qRT-PCR analysis, TAG content, and total FA content are expressed as means ± SDs. Statistical analysis was conducted by one-way ANOVA with the SPSS, version 20.0, program for Windows (SPSS Inc., Values of p ≤ 0.05 were considered statistically significant. The Pearson’s correlation coefficient was evaluated using the Excel Analysis ToolPak (Microsoft, Redmond, WA, USA). Three independent biological replicates were used for each statistical analysis.

Availability of data and materials

The datasets used and/or analyzed during the current study are available from the corresponding author upon reasonable request. Materials will be made available under a material transfer agreement.



Diacylglycerol acyltransferase1-2


Dry weight


Fatty acid


Glyceraldehyde 3-phosphate dehydrogenase


Neomycin phosphotransferase II




Real-time quantitative reverse transcription PCR




Trigalactosyl diacylglycerol1








  1. International Energy Agency. Energy technology perspectives 2017. Accessed 28 Apr 2022.

  2. Field JL, Richard TL, Smithwick EAH, Cai H, Laser MS, LeBauer DS, Long SP, Paustian K, Qin Z, Sheehan JJ, Smith P, Wang MQ, Lynd LR. Robust paths to net greenhouse gas mitigation and negative emissions via advanced biofuels. Proc Natl Acad Sci. 2020;117:21968–77.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Kumar D, Long SP, Arora A, Singh V. Techno-economic feasibility analysis of engineered energycane-based biorefinery co-producing biodiesel and ethanol. GCB Bioenergy. 2021;13:1498–514.

    Article  CAS  Google Scholar 

  4. Durrett TP, Benning C, Ohlrogge J. Plant triacylglycerols as feedstocks for the production of biofuels. Plant J. 2008;54:593–607.

    Article  CAS  PubMed  Google Scholar 

  5. Susmozas A, Martín-Sampedro R, Ibarra D, Eugenio ME, Iglesias R, Manzanares P, Moreno AD. Process strategies for the transition of 1G to advanced bioethanol production. Processes. 2020;8:1310.

    Article  CAS  Google Scholar 

  6. Thelen JJ, Ohlrogge JB. Metabolic engineering of fatty acid biosynthesis in plants. Metab Eng. 2002;4:12–21.

    Article  CAS  PubMed  Google Scholar 

  7. Gurr MI. The biosynthesis of triacylglycerols. In: Stumpf PK, editor. Lipids: structure and function. Amsterdam: Elsevier; 1980. p. 205–48.

    Chapter  Google Scholar 

  8. Ohlrogge J, Chapman K. The seeds of green energy: expanding the contribution of plant oils as biofuels. Biochemist. 2011;33:34–8.

    Article  Google Scholar 

  9. Vanhercke T, Dyer JM, Mullen RT, Kilaru A, Rahman MM, Petrie JR, Green AG, Yurchenko O, Singh SP. Metabolic engineering for enhanced oil in biomass. Prog Lipid Res. 2019;74:103–29.

    Article  CAS  PubMed  Google Scholar 

  10. Zale J, Jung JH, Kim JY, Pathak B, Karan R, Liu H, Chen X, Wu H, Candreva J, Zhai Z, Shanklin J, Altpeter F. Metabolic engineering of sugarcane to accumulate energy-dense triacylglycerols in vegetative biomass. Plant Biotechnol J. 2016;14:661–9.

    Article  CAS  PubMed  Google Scholar 

  11. Sanjaya, Miller R, Durrett TP, Kosma DK, Lydic TA, Muthan B, Koo AJK, Bukhman YV, Reid GE, Howe GA, Ohlrogge J, Benning C. Altered lipid composition and enhanced nutritional value of Arabidopsis leaves following introduction of an algal diacylglycerol acyltransferase 2. Plant Cell. 2013;25:677–93.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Fan J, Zhai Z, Yan C, Xu C. Arabidopsis TRIGALACTOSYLDIACYLGLYCEROL5 interacts with TGD1, TGD2, and TGD4 to facilitate lipid transfer from the endoplasmic reticulum to plastids. Plant Cell. 2015;27:2941–55.

    CAS  PubMed  PubMed Central  Google Scholar 

  13. Roston RL, Gao J, Murcha MW, Whelan J, Benning C. TGD1, -2, and -3 proteins involved in lipid trafficking form ATP-binding cassette (ABC) transporter with multiple substrate-binding proteins. J Biol Chem. 2012;287:21406–15.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Xu C, Fan J, Froehlich JE, Awai K, Benning C. Mutation of the TGD1 chloroplast envelope protein affects phosphatidate metabolism in Arabidopsis. Plant Cell. 2005;17:3094–110.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Lehner R, Kuksis A. Biosynthesis of triacylglycerols. Prog Lipid Res. 1996;35:169–201.

    Article  CAS  PubMed  Google Scholar 

  16. Capuano F, Beaudoin F, Napier JA, Shewry PR. Properties and exploitation of oleosins. Biotechnol Adv. 2007;25:203–6.

    Article  CAS  PubMed  Google Scholar 

  17. Parthibane V, Rajakumari S, Venkateshwari V, Iyappan R, Rajasekharan R. Oleosin is bifunctional enzyme that has both monoacylglycerol acyltransferase and phospholipase activities. J Biol Chem. 2012;287:1946–54.

    Article  CAS  PubMed  Google Scholar 

  18. Winichayakul S, Scott RW, Roldan M, Hatier J-HB, Livingston S, Cookson R, Curran AC, Roberts NJ. In vivo packaging of triacylglycerols enhances Arabidopsis leaf biomass and energy density. Plant Physiol. 2013;162:626–39.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Eastmond PJ. SUGAR-DEPENDENT1 encodes a patatin domain triacylglycerol lipase that initiates storage oil breakdown in germinating Arabidopsis seeds. Plant Cell. 2006;18:665–75.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Fan J, Yan C, Roston R, Shanklin J, Xu C. Arabidopsis lipins, PDAT1 acyltransferase, and SDP1 triacylglycerol lipase synergistically direct fatty acids toward β-oxidation, thereby maintaining membrane lipid homeostasis. Plant Cell. 2014;26:4119–34.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. Kelly AA, van Erp H, Quettier A-L, Shaw E, Menard G, Kurup S, Eastmond PJ. The sugar-dependent1 lipase limits triacylglycerol accumulation in vegetative tissues of Arabidopsis. Plant Physiol. 2013;162:1282–9.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. Slocombe SP, Cornah J, Pinfield-Wells H, Soady K, Zhang Q, Gilday A, Dyer JM, Graham IA. Oil accumulation in leaves directed by modification of fatty acid breakdown and lipid synthesis pathways. Plant Biotechnol J. 2009;7:694–703.

    Article  CAS  PubMed  Google Scholar 

  23. Vanhercke T, El Tahchy A, Liu Q, Zhou XR, Shrestha P, Divi UK, Ral JP, Mansour MP, Nichols PD, James CN, Horn PJ, Chapman KD, Beaudoin F, Ruiz-López N, Larkin PJ, de Feyter RC, Singh SP, Petrie JR. Metabolic engineering of biomass for high energy density: oilseed-like triacylglycerol yields from plant leaves. Plant Biotechnol J. 2014;12:231–9.

    Article  CAS  PubMed  Google Scholar 

  24. Fan J, Yan C, Zhang X, Xu C. Dual role for phospholipid: diacylglycerol acyltransferase: enhancing fatty acid synthesis and diverting fatty acids from membrane lipids to triacylglycerol in Arabidopsis leaves. Plant Cell. 2013;25:3506–18.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Vanhercke T, El Tahchy A, Shrestha P, Zhou X-R, Singh SP, Petrie JR. Synergistic effect of WRI1 and DGAT1 coexpression on triacylglycerol biosynthesis in plants. Febs Lett. 2013;587:364–9.

    Article  CAS  PubMed  Google Scholar 

  26. Vanhercke T, Petrie JR, Singh SP. Energy densification in vegetative biomass through metabolic engineering. Biocatal Agric Biotechnol. 2014;3:75–80.

    Article  Google Scholar 

  27. Yang Y, Munz J, Cass C, Zienkiewicz A, Kong Q, Ma W, Sanjaya, Sedbrook J, Benning C. Ectopic expression of WRINKLED1 affects fatty acid homeostasis in Brachypodium distachyon vegetative tissues. Plant Physiol. 2015;169:1836–47.

    CAS  PubMed  PubMed Central  Google Scholar 

  28. Parajuli S, Kannan B, Karan R, Sanahuja G, Liu H, Garcia-Ruiz E, Kumar D, Singh V, Zhao H, Long S, Shanklin J, Altpeter F. Towards oilcane: engineering hyperaccumulation of triacylglycerol into sugarcane stems. GCB Bioenergy. 2020;12:476–90.

    Article  CAS  Google Scholar 

  29. Alameldin H, Izadi-Darbandi A, Smith SA, Balan V, Jones AD, Sticklen M. Production of seed-like storage lipids and increase in oil bodies in corn (maize; Zea mays L.) vegetative biomass. Ind Crops Prod. 2017;108:526–34.

    Article  CAS  Google Scholar 

  30. Vanhercke T, Belide S, Taylor MC, El Tahchy A, Okada S, Rolland V, Liu Q, Mitchell M, Shrestha P, Venables I, Ma L, Blundell C, Mathew A, Ziolkowski L, Niesner N, Hussain D, Dong B, Liu G, Godwin ID, Lee J, Rug M, Zhou XR, Singh SP, Petrie JR. Up-regulation of lipid biosynthesis increases the oil content in leaves of Sorghum bicolor. Plant Biotechnol J. 2019;17:220–32.

    Article  CAS  PubMed  Google Scholar 

  31. Beechey-Gradwell Z, Cooney L, Winichayakul S, Andrews M, Hea SY, Crowther T, Roberts N. Storing carbon in leaf lipid sinks enhances perennial ryegrass carbon capture especially under high N and elevated CO2. J Exp Bot. 2020;71:2351–61.

    Article  CAS  PubMed  Google Scholar 

  32. Matsuoka S, Kennedy AJ, Santos EGD, Tomazela AL, Rubio LCS. Energy cane: Its concept, development, characteristics, and prospects. Adv Bot. 2014;2014:1–13.

    Article  Google Scholar 

  33. Alexander AG. The energy cane alternative. Amsterdam: Elsevier; 1985.

    Google Scholar 

  34. Carvalho-Netto OV, Bressiani JA, Soriano HL, Fiori CS, Santos JM, Barbosa GV, Xavier MA, Landell MG, Pereira GA. The potential of the energy cane as the main biomass crop for the cellulosic industry. Chem Biol Technol Agric. 2014;1:1–8.

    Article  CAS  Google Scholar 

  35. Diniz AL, Ferreira SS, Ten-Caten F, Margarido GR, Dos Santos JM, Barbosa GVS, Carneiro MS, Souza GM. Genomic resources for energy cane breeding in the post genomics era. Comput Struct Biotechnol J. 2019;17:1404–14.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  36. Richard TL. Challenges in scaling up biofuels infrastructure. Science. 2010;329:793–6.

    Article  CAS  PubMed  Google Scholar 

  37. Fouad WM, Hao W, Xiong Y, Steeves C, Sandhu SK, Altpeter F. Generation of transgenic energy cane plants with integration of minimal transgene expression cassette. Curr Pharm Biotechnol. 2015;16:407–13.

    Article  CAS  PubMed  Google Scholar 

  38. Ramasamy M, Mora V, Damaj MB, Padilla CS, Ramos N, Rossi D, Solís-Gracia N, Vargas-Bautista C, Irigoyen S, DaSilva JA, Mirkov TE. A biolistic-based genetic transformation system applicable to a broad-range of sugarcane and energycane varieties. GM Crops Food. 2018;9:211–27.

    Article  PubMed  PubMed Central  Google Scholar 

  39. Padilla CS, Damaj MB, Yang Z-N, Molina J, Berquist BR, White EL, Solís-Gracia N, Da Silva J, Mandadi KK. High-level production of recombinant snowdrop lectin in sugarcane and energy cane. Front Bioeng Biotechnol. 2020;8:977.

    Article  PubMed  PubMed Central  Google Scholar 

  40. Eid A, Mohan C, Sanchez S, Wang D, Altpeter F. Multiallelic, targeted mutagenesis of magnesium chelatase with CRISPR/Cas9 provides a rapidly scorable phenotype in highly polyploid sugarcane. Front Genome Ed. 2021;3:654996.

    Article  PubMed  PubMed Central  Google Scholar 

  41. Kannan B, Jung JH, Moxley GW, Lee SM, Altpeter F. TALEN-mediated targeted mutagenesis of more than 100 COMT copies/alleles in highly polyploid sugarcane improves saccharification efficiency without compromising biomass yield. Plant Biotechnol J. 2018;16:856–66.

    Article  CAS  PubMed  Google Scholar 

  42. Oz MT, Altpeter A, Karan R, Merotto A, Altpeter F. CRISPR/Cas9-mediated multi-allelic gene targeting in sugarcane confers herbicide tolerance. Front Genome Ed. 2021;3:673566.

    Article  PubMed  PubMed Central  Google Scholar 

  43. Zhao Y, Karan R, Altpeter F. Error-free recombination in sugarcane mediated by only 30 nucleotides of homology and CRISPR/Cas9 induced DNA breaks or Cre-recombinase. Biotechnol J. 2021;16:2000650.

    Article  CAS  Google Scholar 

  44. Mudge SR, Basnayake SW, Moyle RL, Osabe K, Graham MW, Morgan TE, Birch RG. Mature-stem expression of a silencing-resistant sucrose isomerase gene drives isomaltulose accumulation to high levels in sugarcane. Plant Biotechnol J. 2013;11:502–9.

    Article  CAS  PubMed  Google Scholar 

  45. Wang J, Li Y, Wai CM, Beuchat G, Chen L-Q. Identification and analysis of stem-specific promoters from sugarcane and energy cane for oil accumulation in their stems. GCB Bioenergy. 2021;13:1515–27.

    Article  CAS  Google Scholar 

  46. Mitchell MC, Pritchard J, Okada S, Zhang J, Venables I, Vanhercke T, Ral J-P. Increasing growth and yield by altering carbon metabolism in a transgenic leaf oil crop. Plant Biotechnol J. 2020;18:2042–52.

    Article  CAS  PubMed Central  Google Scholar 

  47. Tai PYP, Shine JM Jr, Glaz B, Deren CW, Miller JD, Comstock JC. Registration of ‘CP 82–1592’ Sugarcane. Crop Sci. 1991;31:1706–7.

    Google Scholar 

  48. Murray MG, Thompson WF. Rapid isolation of high molecular weight plant DNA. Nucleic Acids Res. 1980;8:4321–6.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  49. Kuijper J. De groei van bladschijf, bladscheede em stengel van het suikerriet. Arch Suikerind Ned-Indie. 1915;23:528–56.

    Google Scholar 

  50. Iskandar HM, Simpson RS, Casu RE, Bonnett GD, Maclean DJ, Manners JM. Comparison of reference genes for quantitative real-time polymerase chain reaction analysis of gene expression in sugarcane. Plant Mol Biol Rep. 2004;22:325–37.

    Article  CAS  Google Scholar 

  51. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods. 2001;25:402–8.

    Article  CAS  PubMed  Google Scholar 

Download references


The authors would like to thank Dr. Hardev Sandhu, Everglades Research and Education Center, UF-IFAS, for providing energycane tops and Sun Gro Horticulture, Apopka, FL, for the donation of potting mix.


This research was funded by the Biological and Environmental Research (BER) program, U.S. Department of Energy, under Award Number DE-SC0018254. The views and opinions of authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof. This work was also supported by the USDA National Institute of Food and Agriculture, Hatch project 1020425.

Author information

Authors and Affiliations



FA and JS conceived and designed the experiments. FA designed the recombinant DNA constructs. GL and VDC carried out the tissue culture and transformation of energycane, grew the transgenic plants in the greenhouse, and sampled for TAG, FA, and qRT-PCR analysis. GL, VDC, and BK carried out qRT-PCR analysis. HL conducted the analyses for TAG, and the total content and composition of fatty acids. VDC completed the statistical analysis. GL, VDC, and FA wrote the manuscript. All authors read and approved the final manuscript.

Corresponding authors

Correspondence to John Shanklin or Fredy Altpeter.

Ethics declarations

Ethics approval and consent to participate

The research on transgenic energycane was carried out in compliance with relevant institutional and national guidelines.

Consent for publication

Not applicable.

Competing interests

The authors have no competing interests to declare.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Additional file 1. Table S1

. List of primers used for gene expression analysis; Table S2. Summary of PCR analysis of transgenic lines; Table S3. Summary of TAG content and expression/suppression of (trans)genes in leaves of transgenic sugarcane; Table S4. Correlation of total FA content with TAG content in transgenic energycane; Figure S1. PCR analysis of transgenic plants.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Luo, G., Cao, V.D., Kannan, B. et al. Metabolic engineering of energycane to hyperaccumulate lipids in vegetative biomass. BMC Biotechnol 22, 24 (2022).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: