Lessons learned from metabolic engineering of cyanogenic glucosides
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- Morant, A.V., Jørgensen, K., Jørgensen, B. et al. Metabolomics (2007) 3: 383. doi:10.1007/s11306-007-0079-x
Plants produce a plethora of secondary metabolites which constitute a wealth of potential pharmaceuticals, pro-vitamins, flavours, fragrances, colorants and toxins as well as a source of natural pesticides. Many of these valuable compounds are only synthesized in exotic plant species or in concentrations too low to facilitate commercialization. In some cases their presence constitutes a health hazard and renders the crops unsuitable for consumption. Metabolic engineering is a powerful tool to alter and ameliorate the secondary metabolite composition of crop plants and gain new desired traits. The interplay of a multitude of biosynthetic pathways and the possibility of metabolic cross-talk combined with an incomplete understanding of the regulation of these pathways, explain why metabolic engineering of plant secondary metabolism is still in its infancy and subject to much trial and error. Cyanogenic glucosides are ancient defense compounds that release toxic HCN upon tissue disruption caused e.g. by chewing insects. The committed steps of the cyanogenic glucoside biosynthetic pathway are encoded by three genes. This unique genetic simplicity and the availability of the corresponding cDNAs have given cyanogenic glucosides pioneering status in metabolic engineering of plant secondary metabolism. In this review, lessons learned from metabolic engineering of cyanogenic glucosides in Arabidopsis thaliana (thale cress), Nicotiana tabacum cv Xanthi (tobacco), Manihot esculenta Crantz (cassava) and Lotus japonicus (bird’s foot trefoil) are presented. The importance of metabolic channelling of toxic intermediates as mediated by metabolon formation in avoiding unintended metabolic cross-talk and unwanted pleiotropic effects is emphasized. Likewise, the potential of metabolic engineering of plant secondary metabolism as a tool to elucidate, for example, the impact of secondary metabolites on plant–insect interactions is demonstrated.
KeywordsCyanogenic glucosides Metabolic engineering RNAi Transgene silencing 5′-Azacytidine
Genetic engineering provides a valuable approach to alter and improve metabolite composition in crop plants to generate robust plants with improved traits of interest to consumers and producers. Plants can be genetically engineered to have higher nutritional value or tailor-made resistance against pathogens and herbivores. Genetic engineering also holds promise for economically favourable production platforms for bio-pharmaceuticals, essential oils, colorants, flavours and fragrances. Examples of genetic modification of crops to improve nutritional quality include “Golden Rice” with increased pro-vitamin A content (Ye et al. 2000), oil crops accumulating essential very long chain polyunsaturated fatty acids (Abbadi et al. 2004; Wu et al. 2005), potatoes that accumulate storage proteins with increased levels of essential amino acids (Chakraborty et al. 2000), iron and zinc enriched rice (Vasconcelos et al. 2003) and low allergen soybean (Herman et al. 2003). Plant metabolism is highly complex, and predictive metabolic engineering is often hampered by a lack of detailed knowledge about metabolic cross-talk and regulation of metabolic grids. A major challenge in metabolic engineering is to design and construct plants with a limited number of unintended side effects and to reduce the number of unexpected results by enhancing our ability to carry out in silico prediction of metabolic responses to alterations in biosynthetic pathways.
Of particular interest is the ability to engineer plants with new or altered levels of secondary metabolites often referred to as natural products. Plants have the capacity to synthesize a vast range of secondary metabolites making plants the organic chemist par excellence in nature. The impact of secondary metabolites in the successful adaption of plant species cannot be underestimated as these highly sophisticated small metabolites have evolved during millions of years of selection during speciation. They are important players in adaptation to abiotic and biotic stresses such as acclimation and plant–insect and plant–microbe interactions, as they provide the chemical signals that enable plants to deter herbivores and pests, attract pollinators, communicate with other plants and constantly adapt to climatic changes. Through time, man has relied upon and exploited the use of plant secondary metabolites as flavours, scents, poisons, natural pesticides and pharmaceuticals. Not only are secondary metabolites the active components in traditional herbal medicines, they are often the origin and/or the precursor of most of today’s medicine (Morant et al. 2003). The exploitation of plant secondary metabolites is often hampered by their accumulation in too low amounts or by the fact that their occurrence is restricted to a single or few exotic plants not suited for commercialization. In addition, a lack of knowledge of the enzymatic steps and, in particular, the unavailability of the underlying genes encoding the requisite enzymes present major limitation to the exploitation of plants as green factories for production of desired secondary metabolites (Kutchan 2005).
Cyanogenic glucosides constitute a limited number of amino acid derived secondary metabolites found throughout the plant kingdom (Bak et al. 2006). What makes cyanogenic glucosides interesting from a metabolic engineering point of view is that this group of compounds is thought to play an important role in the ability of plants to combat pests and diseases. In addition, the entire pathway has been elucidated and the cDNAs encoding the three enzymes that catalyze the committed steps have been isolated. The availability of the cDNAs and the unique genetic simplicity have given the cyanogenic glucoside pathway its pioneering status in metabolic engineering.
Transfer of the entire dhurrin pathway from S. bicolor into a distantly related plant species using genetic engineering has provided a powerful tool to study the impact of dhurrin as a defense compound (Tattersall et al. 2001), because this offers the possibility to eliminate the impact of natural variation with respect to metabolite composition and morphology which is otherwise often encountered when ecotypes and mutants are used to study plant–insect interactions. Expression of enzymes from the dhurrin pathway in Arabidopsis thaliana (thale cress) offers the opportunity to study metabolic cross-talk between the structurally related cyanogenic glucoside and glucosinolate pathways, as well as the impact of dhurrin and cyanogenesis on insects that specifically feed on cruciferous plants such as the cruciferous feeding specialist flea beetle Phyllotreta nemorum (Tattersall et al. 2001). A. thaliana is easy to transform and its entire genome has been sequenced which makes it an obvious choice for genetic engineering. As a cruciferous plant, A. thaliana does not synthesize cyanogenic glucosides but produces glucosinolates, a group of amino acid derived phytoanticipins related to, but not naturally co-occurring with, cyanogenic glucosides (Fig. 1C; Halkier and Gershenzon 2006). While cyanogenic glucosides are synthesized from tyrosine, phenylalanine, leucine, isoleucine, valine and 2-cyclopentenylglycine, glucosinolates are derived from methionine, alanine, isoleucine, valine, leucine, tryptophan, tyrosine and phenylalanine by a biosynthetic pathway whose initial reaction steps are equivalent to those of the cyanogenic glucoside pathway (Bak et al. 1998b). In both pathways, a multifunctional cytochrome P450 belonging to the CYP79 family catalyzes the conversion of the precursor amino acid or the chain elongated amino acid into the corresponding oxime which is further metabolized by a second cytochrome P450 belonging to either the CYP71 or CYP83 family (Fig. 1B, C; Bak et al. 2006; Halkier and Gershenzon 2006).
The leguminous plant Lotus japonicus (bird’s foot trefoil) accumulates the aliphatic cyanogenic glucosides linamarin and lotaustralin derived from valine and isoleucine, respectively, and the isoleucine-derived cyanoalkenyl glucosides, rhodiocyanoside A and D (Forslund et al. 2004). Rhodiocyanosides constitute a class of glucosides that are related to the cyanogenic glucosides, but whose biological function is currently not understood. While rhodiocyanoside A and D are degraded by the same β-glucosidases that degrade the cyanogenic glucosides (our unpublished results), hydrolysis is not accompanied by release of HCN, as rhodiocyanosides are not derived from α-hydroxynitriles (cyanohydrins). While the ability to produce linamarin and lotaustralin is widespread in Lotus species, the ability to produce rhodiocyanosides appears to be limited to L. japonicus (Zagrobelny et al. 2007 and our unpublished results).
Flea beetles have co-evolved with cruciferous plants and have adapted to the presence of glucosinolates in their diet. In a similar manner, Zygaena species (burnet moths) have co-evolved with Lotus species. Zygaena larvae feed on Fabaceous plants including Lotus species and sequester the cyanogenic glucosides for use in their own defense against predators. In the absence of sufficient amounts of cyanogenic glucosides in their dietary plants, the larvae possess the ability to de novo synthesize the very same cyanogenic glucosides, linamarin and lotaustralin, as present in Lotus species, albeit with a resultant concomitant reduction in growth rate (Zagrobelny et al. 2004). Metabolic engineering of L. japonicus to obtain plants with altered cyanogenic glucoside profiles, i.e. by introduction of novel cyanogenic glucosides such as dhurrin, or silencing of the endogenous cyanogenic glucoside pathway to obtain plants depleted of cyanogenic glucosides and rhodiocyanosides, thus provides a unique opportunity to elucidate the impact of cyanogenic glucosides on plant–insect interactions using the unique Zygaena—Lotus system.
Similar to Lotus species, the key stable crop Manihot esculenta Crantz (cassava) contains the cyanogenic glucosides linamarin and lotaustralin. M. esculenta is the world’s most important tropical root crop and serves as a famine-reserve in the third world, especially in Africa (Nweke et al. 2002). A major nutritive drawback is the accumulation of up to 1.5 g/kg dry weight of linamarin and lotaustralin in the M. esculenta tubers (Bokanga 1994). Consequently it is of great interest to develop M. esculenta with acyanogenic tubers to provide a healthier diet for millions of people in the third world.
In this review, we report the lessons learned from studies on engineering the dhurrin biosynthetic pathway from S. bicolor into A. thaliana and L. japonicus and from silencing cyanogenic glucoside biosynthesis in M. esculenta and L. japonicus. We conclude that predictive metabolic engineering requires not only understanding of the metabolic pathways of the plant in question and the engineered pathway in particular, but also of transport and accumulation of the novel product as well as the ability of the plant to accommodate the transgenes and their encoded proteins.
1.1 Engineering metabolic cross-talk between the glucosinolate and cyanogenic glucoside pathways in A. thaliana changes the glucosinolate profile
The total level of glucosinolates in A. thaliana lines that express S. bicolor CYP79A1 is four times higher compared to wild type, i.e. p-hydroxybenzylglucosinolate accounts for ∼75% of the total amount of glucosinolates (Bak et al. 1999). In spite of this, two species of flea beetles, P. nemorum and P. cruciferae did not discriminate between A. thaliana 35S::CYP79A1 and wild type in free choice feeding experiments (Nielsen et al. 2001). This demonstrates that neither a significant change in total glucosinolate content nor glucosinolate profile is important for the ability of flea beetles to recognize and feed on A. thaliana.
In a similar series of experiments, S. bicolor CYP79A1 was expressed in transgenic Nicotiana tabacum cv Xanthi (tobacco) plants (Bak et al. 2000). The transgenic N. tabacum plants are reduced in height, produce a very limited number of seeds and accumulate metabolites derived from detoxification of p-hydroxyphenylacetaldoxime (our unpublished results).
Subsequent to the production of the transgenic A. thaliana plants producing a new tyrosine-derived glucosinolate, A. thaliana plants have been engineered to accumulate high amounts (up to 35% of total glucosinolate content in mature rosette leaves) of valine- and isoleucine-derived glucosinolates (Mikkelsen and Halkier 2003). This was achieved by expression of M. esculenta CYP79D2, the gene encoding the enzyme that catalyzes the conversion of valine and isoleucine to the corresponding aldoximes in the cyanogenic glucoside pathway in M. esculenta (Andersen et al. 2000). These results confirm that the substrate specificity of the CYP79 enzymes is a major determinant of the glucosinolate profile and substantiate the broad substrate specificity of the post oxime enzymes in the glucosinolate pathway.
1.2 Metabolic engineering of dhurrin biosynthesis in A. thaliana: Efficient channelling of biosynthetic intermediates and resistance to flea beetles
A major break through in metabolic engineering of secondary metabolites in plants was the ability to introduce the cyanogenic glucoside dhurrin into A. thaliana plants with marginal impact on visual plant phenotype, metabolome and transcriptome (Kristensen et al. 2005; Kutchan 2005; Memelink 2005; Tattersall et al. 2001). The transgenic A. thaliana lines producing high levels (up to 4% dry weight) of the tyrosine-derived dhurrin was generated by two consecutive transformation events. First the two cytochromes P450, CYP79A1 and CYP71E1, were introduced via a single construct, p2x (Fig. 1D; Bak et al. 2000). These transgenic A. thaliana lines, designated 2x, appeared stressed and stunted in growth due to accumulation of toxic dhurrin intermediates and derivatives thereof (Fig. 2, panel 2x; Bak et al. 2000; Kristensen et al. 2005; Tattersall et al. 2001). In contrast, A. thaliana plants transformed with S. bicolor CYP79A1 did not display a visual phenotype. Co-transformation of CYP79A1 with CYP71E1 generates a new sink for p-hydroxyacetaldoxime metabolism and efficiently prevents redirection of the majority of the generated p-hydroxyphenylacetaldoxime into the glucosinolate pathway. Accordingly, the A. thaliana 2x lines preferentially produce the unstable cyanohydrin p-hydroxymandelonitrile which decomposes into primarily p-hydroxybenzoic acid via p-hydroxybenzaldehyde (Bak et al. 2000; Kristensen et al. 2005; Tattersall et al. 2001). In the 2x lines, the p-hydroxybenzoic acid is glucosylated and accumulates as p-hydroxybenzoylglucoside (Fig. 2, panel 2x). Unexpectedly, the UV protectants sinapoyl malate and sinapoyl glucose were decreased in the A. thaliana 2x lines (Fig. 2; Kristensen et al. 2005).
To complete the dhurrin biosynthetic pathway, A. thaliana 2x lines were re-transformed with pGT (Fig. 1D), thereby introducing UGT85B1, the UDP-glucosyl transferase that catalyzes the final step in biosynthesis of dhurrin in S. bicolor (Jones et al. 1999). Upon introduction of UGT85B1 (Fig. 1B, C), the visual phenotype was restored to wild type and up to 4% dry weight dhurrin accumulated. As a consequence, levels of sinapoyl malate and sinapoyl glucose were restored to wild type, and the detoxification products observed in the recipient 2x lines were no longer detectable (Fig. 2, panel 3x; Kristensen et al. 2005; Tattersall et al. 2001). Notably, p-hydroxybenzylglucosinolate was not detected in the A. thaliana 3x lines, indicating that in the presence of the entire set of enzymes catalyzing the dhurrin pathway, the glucosinolate pathway is no longer able to compete for the p-hydroxyphenylacetaldoxime generated by S. bicolor CYP79A1. A likely explanation is that when S. bicolor CYP79A1, CYP71E1, and UGT85B1 are co-expressed, they form a tight metabolon that effectively channels tyrosine to dhurrin (Jørgensen et al. 2005b; Møller and Conn 1980) and simultaneously prevents S. bicolor CYP79A1 from interacting with the post oxime enzymes in the glucosinolate pathway. These results prove that it is possible to engineer transgenic plants that produce significant amounts of a novel secondary metabolite and yet adhere to the principle of substantial equivalence (Kristensen et al. 2005).
Despite the high levels of dhurrin, the A. thaliana 3x lines are not highly cyanogenic, i.e. they are only able to slowly degrade dhurrin. This reflects the lack of a specific β-glucosidase catalyzing immediate hydrolysis of dhurrin. Consequently, tissue damage results in a cyanide fizz rather than a cyanide bomb (Tattersall et al. 2001). Whereas the cruciferous specialist flea beetle P. nemorum did not discriminate between the A. thaliana 1x lines that accumulate high levels of p-hydroxybenzylglucosinolate and wild type leaves (Nielsen et al. 2001), a significant deterrent effect in choice tests between A. thaliana 3x and wild type was observed. The flea beetles consumed up to 80% less of the A. thaliana 3x leaf material as compared to wild type (Tattersall et al. 2001). Similarly, the majority of flea beetle larvae died when fed the dhurrin containing 3x lines (Tattersall et al. 2001). Experiments in which the flea beetles were starved for 2 days and subsequently fed dhurrin producing A. thaliana 3x plants in non-choice experiments, revealed that the flea beetles did consume leaf material from the cyanogenic plants, but that this resulted in transient paralysis in their legs (our unpublished data). These results unambiguously confirmed that cyanogenic glucosides may confer resistance to herbivores. The results also served to illustrate the inherent ability of animals to detoxify HCN (Zagrobelny et al. 2004) in that paralysis was only transient.
1.3 Expression of the dhurrin biosynthetic pathway in L. japonicus
Sorghum bicolor accumulates the aromatic cyanogenic glucoside dhurrin derived from tyrosine while L. japonicus accumulates the aliphatic cyanogenic glucosides linamarin and lotaustralin derived from valine and isoleucine, respectively. Expression of S. bicolor CYP79A1, CYP71E1 and UGT85B1 either separately or in concert in L. japonicus would facilitate studies of the flexibility of the cyanogenic glucoside pathway in a cyanogenic plant. This would yield valuable information on the ability of the enzymes of one pathway to interact and enter into a metabolon with enzymes of the parallel pathway, on the substrate specificity of the post oxime enzymes and on the capability of L. japonicus to host an entire heterologous pathway and synthesize and store a new cyanogenic glucoside. In addition such plants would facilitate a study of the impact of the total cyanogenic glucoside content as well as the cyanogenic glucoside profile on the interactions between L. japonicus and Zygaena species. Accordingly the dhurrin pathway was introduced into L. japonicus. To achieve this, three approaches were pursued. The first approach was analogous to the introduction of the tyrosine-derived p-hydroxybenzylglucosinolate in A. thaliana taking advantage of an expected relatively broad substrate specificity of the post oxime enzymes in the endogenous L. japonicus cyanogenic glucoside pathway. Previously, microsomes prepared from L. japonicus leaves have been shown to convert p-hydroxyphenylacetaldoxime into the corresponding cyanohydrin, p-hydroxymandelonitrile (Forslund et al. 2004), thus demonstrating that the post oxime enzymes in L. japonicus are able to metabolize p-hydroxyphenylacetaldoxime. Free oximes are generally known to be toxic to the plant (Bak et al. 1999; Grootwassink et al. 1990; Hemm et al. 2003) and thus the ability of endogenous enzymes to metabolize p-hydroxyphenylacetaldoxime is a prerequisite for successful expression of S. bicolorCYP79A1 in L. japonicus. Accordingly, L. japonicus were transformed with construct p1x (Fig. 1D) to introduce S. bicolor CYP79A1. However, no transformants were obtained that expressed S. bicolor CYP79A1. In L. japonicus, the cyanogenic glucoside pathway was subsequently shown not be expressed in the callus phase (our unpublished data) as also observed in M. esculenta (Joseph et al. 1999). Accordingly, successfully transformed cells or calli would accumulate toxic p-hydroxyphenylacetaldoxime and as a consequence most likely die of intoxication or silence the transgene. In retrospect, this approach thus appears suboptimal.
The second approach was based on the ability to transform the three cDNAs encoding the enzymes of the dhurrin pathway into A. thaliana (Tattersall et al. 2001). Accordingly, experiments were set up to initially transform L. japonicus with the p2x construct (Fig. 1D) with a planned re-transformation with pGT. However, it was not possible to regenerate shoots from the L. japonicus 2x explants. The fact that A. thaliana but not L. japonicus may be successfully transformed with the p2x construct is most probably due to a combination of factors that each imposes a negative selection against p2x transformed L. japonicus explants. A major difference relates to the transformation procedure employed. L. japonicus transformation requires an extended callus phase (Handberg and Stougaard 1992), in which the plant cells remain in an undifferentiated state, whereas A. thaliana is transformed by simply dipping developing flowers into a solution of Agrobacterium and subsequently harvesting and selecting the transformed seeds (Clough and Bent 1998). Moreover, A. thaliana is able to detoxify a proportion of the tyrosine-derived oxime by redirection into the glucosinolate pathway. Finally, L. japonicus 2x explants most likely do not possess the physiological machinery to handle the toxic compounds produced by CYP79A1 and CYP71E1, and are probably subjected to cyanide intoxication. N. tabacum and Vitis vinifera L. (grapevine), two species that likewise require a callus phase as part of the transformation procedure have been successfully transformed with p2x, and expression of CYP79A1 and CYP71E1 obtained (Bak et al. 2000; Franks et al. 2006). A major difference between these three species is that the callus phase in the course of N. tabacum transformation is significantly shorter (∼1 month) than the callus phase required for L. japonicus transformation (3–6 months), while V. vinifera transformation involves an intermediate 2–3 months callus phase (Iocco et al. 2001). In addition, out of 35 N. tabacum 2x transformants, only ten lines produced detectable amounts of CYP79A1 and CYP71E1 enzyme activity and with significantly lower enzyme activity compared to A. thaliana 2x (Bak et al. 2000). Likewise, only 2 out of 19 kanamycin resistant V. vinifera 2x transformants expressed detectable CYP79A1 and CYP71E1 (Franks et al. 2006). These results suggest that transformants which expressed high levels and/or highly active CYP79A1 and CYP71E1 are selected against during the callus phase or during regeneration.
1.3.1 Transgene silencing and re-activation of dhurrin biosynthesis by 5′-azacytidine treatment
The ratio of dhurrin producing L. japonicus 3x progeny observed when germinated in the presence of azaC corresponds to an expected 9/16 ratio based on simple Mendelian segregation of the two T-DNAs derived from p2x and pGT in a heterozygous parent plant. Re-activation of dhurrin production proved to be transient as silencing recurred in all plants examined at the latest 6 weeks after germination in the presence of azaC. This is most likely linked to the instability of azaC as well as the dilution resulting from repeated cell division. L. japonicus is thus able to recognize and inactivate the transgenes once azaC levels have been diluted below a certain threshold. Re-activation of silenced transgenes in rice by azaC treatment followed by new onset of silencing was similarly observed in the study by Kumpatla and Hall (1998).
Transcriptional gene silencing, however, does not exclude PTGS as a determining factor in silencing of the dhurrin pathway in L. japonicus 3x. To examine a possible additional effect of PTGS, L. japonicus 3x could be transformed with a viral suppressor of PTGS such as the p19 protein from tomato bushy stunt virus. The p19 protein is well known to enhance expression of transgenes in N. tabacum (Voinnet et al. 2003). Studying the effect of p19 in the presence and absence of azaC might reveal any additive effect of PTGS in the observed silencing. We have not pursued this possibility as it would necessitate a transformation round and a subsequent laborious screening for lines that successfully express all three cDNA constructs in a population segregating for three traits.
In addition to dhurrin, two new tyrosine-derived compounds were detected upon administration of radio-labelled tyrosine to detached leaves of L. japonicus 3x germinated in the presence of azaC (Fig. 3B, lane 3). In comparison, dhurrin was the only new metabolite detected in A. thaliana 3x leaves [Figs. 2 (panel 3x), 3B (lane 4)]. LC-MS analysis revealed that azaC treated L. japonicus 3x produced three new unknown compounds in addition to dhurrin (m/z 311) [Fig. 4, (c1, c2 and c3)] with m/z of 300, 397 and 386, respectively. The UV spectra recorded indicated that c1, c2 and c3 are aromatic and derived from tyrosine (results not shown) in agreement with the results obtained when radio-labelled tyrosine was administered (Fig. 3B). As none of the three unknown compounds were detected in extracts from L. japonicus 3x germinated in the absence of azaC, nor in wild type L. japonicus germinated with or without azaC (Fig. 4), this strongly indicates that c1, c2 and c3 are related to the inserted dhurrin pathway.
1.3.2 Why does L. japonicus silence the dhurrin pathway?
Transgene silencing was not observed in L. japonicus 35S::CYP79D2 (Forslund et al. 2004) nor in L. japonicus 35S::UGT85B1 primary transformants. These L. japonicus lines each held two copies of the CaMV 35S promoter. The most likely cause of the silencing observed in L. japonicus 3x is homology-dependent TGS due to the presence of at least five copies of the CaMV 35S promoter that drive the expression of the transgenes and the selective marker genes (Fig. 1D). This explanation is supported by the fact that S. bicolor UGT85B1 is highly active in this L. japonicus GT line but becomes silenced upon re-transformation with S. bicolor CYP79A1 and CYP71E1 (Fig. 3A, lanes 1, 2). In agreement with this hypothesis, strong S. bicolor UGT85B1 activity was observed in false positive transformants (L. japonicus GT lines that escaped kanamycin selection) which indicates that loss of GT activity was caused by introduction of S. bicolor CYP79A1 and CYP71E1 and not by passing through a second callus phase. In the attempt to introduce dhurrin into V. vinifera hairy roots, accumulation of S. bicolor CYP79A1 and CYP71E1 transcripts was detected in V. vinifera 2x (Franks et al. 2006). Upon re-transformation with S. bicolor UGT85B1, some transgenic lines produced dhurrin while in others, all three transgenes were silenced. Hence re-transformation with S. bicolor UGT85B1 silenced the previously expressed S. bicolor CYP79A1 and CYP71E1. In analogy with our results, the authors suggested that the lack of transgene expression was a result of homology-dependent TGS caused by the presence of multiple CaMV 35S promoters (Franks et al. 2006).
In conclusion, the main differences between A. thaliana 3x and L. japonicus 3x transgenic lines are that A. thaliana 3x was transformed by the floral dip method (Clough and Bent 1998) and initially with p2x followed by pGT, while L. japonicus transformation involved an extended callus phase (Handberg and Stougaard 1992) and initial transformation with pGT followed by re-transformation with p2x. The A. thaliana 3x transgenic lines studied by Kristensen et al. (2005) were homozygous for the transgenes and hence held ten copies of the CaMV 35S promoter. As transformation of A. thaliana with CYP79A1 and CYP71E1 results in a severe phenotype, the 2x transformants are likely to have been selected for low amounts or reduced activity of the two cytochromes P450. In agreement with this, DNA microarray data showed that CYP79A1, CYP71E1 as well as UGT85B1 were expressed at low levels in spite of being expressed under control of the CaMV 35S promoter (Kristensen et al. 2005). These data indicate that even low concentrations of the transcripts encoding the dhurrin biosynthetic enzymes are sufficient for production of high amounts of dhurrin. In contrast, the L. japonicus GT line chosen for re-transformation with p2x had high GT activity (Fig. 3A, lane 1). These results indicate that low enzyme activity might be preferable for production of dhurrin in a heterologous plant species and that choice of a low UGT85B1 activity L. japonicus line for p2x transformation line might have been preferable in order to avoid transgene silencing.
1.4 Changing the cyanogenic glucoside profile in transgenic M. esculenta and L. japonicus
1.4.1 The quest for acyanogenic M. esculenta
The low cyanogen (<25% of wild type cyanide potential) M. esculenta RNAi transformants had long, slender stems with long internodes when grown in vitro. When the low cyanogen lines where moved to high nitrogen media or to soil, the phenotype was complemented (Jørgensen et al. 2005a, b). The restored visual phenotype was not due to increased accumulation of cyanogenic glucosides, given that ∼90% of the soil grown RNAi M. esculenta lines retained the >99% reduction in leaf cyanogenic glucoside content as compared to wild type. Apart from a direct involvement in plant defense, cyanogenic glucosides have also been claimed to act as nitrogen storage compounds (Selmar et al. 1988). The phenotypes observed when grown in low nitrogen media might be a reflection of perturbed nitrogen homeostasis or even that cyanogenic glucosides act as signalling compounds that can affect enzyme activities or gene expression involved in plant development. Alternatively, the RNAi construct might have targeted other transcripts in addition to CYP79D1 and CYP79D2 affecting in vitro growth. Future experiments using M. esculenta oligonucleotide DNA micro arrays might help to elucidate the impact of linamarin and lotaustralin on M. esculenta fitness and development.
In the M. esculenta RNAi lines, the cyanogenic glucoside content in tubers varied from 8% to more than 200% of wild type levels in spite of the <1% cyanogenic glucoside content observed in the leaves (Jørgensen et al. 2005a). In M. esculenta, the cyanogenic glucosides are primarily synthesized in leaves and transported to the tubers (Jørgensen et al. 2005a). The accumulation of high amounts of cyanogenic glucosides in tubers from plants with almost acyanogenic leaves indicates a very efficient transport of cyanogenic glucosides from leaves to tubers, and supports that de novo synthesis also takes place in roots of M. esculenta (Du et al. 1995). In addition, a reduced rate of catabolism of cyanogenic glucosides could contribute to the accumulation of cyanogenic glucosides in the tubers of otherwise acyanogenic M. esculenta.
The CaMV 35S promoter is generally regarded as a constitutive, highly active promoter that is active in most plant cell types though numerous plant species dependent exceptions have been reported (e.g. Samac et al. 2004). In M. esculenta, the CaMV35S promoter is known to have reduced activity in the root cells where the CYP71E1 orthologue is expressed (Zhang et al. 2003). Accordingly, a promoter that specifically drives expression at the cellular sites of cyanogenic glucoside biosynthesis is desirable to obtain M. esculenta with acyanogenic tubers. Work is in progress to engineer a CYP79D1/CYP79D2 RNAi construct under control of an endogenous promoter that specifically controls expression of cyanogenic glucoside biosynthesis to facilitate a more targeted silencing. With respect to the use of M. esculenta as a source of food for humans, an optimal transgenic M. esculenta plant would most likely be a plant which produces acyanogenic tubers for consumption while retaining at least a medium cyanide potential in the aerial parts to protect against pests. Engineering of such a M. esculenta variety would require additional knowledge on the genes controlling transport of cyanogenic glucosides, their biosynthesis in roots and possibly the activity of cyanogenic glucoside degrading β-glucosidases.
1.4.2 CYP79D3 and CYP79D4 as targets in the production of L. japonicus with reduced cyanogenic glucoside and rhodiocyanoside content
As an alternative to RNAi technology, isolation of Targeted Induced Local Lesions IN Genomes (TILLING) mutants of L. japonicus is now a possibility with the generation of an L. japonicus TILLING collection (http://www.lotusjaponicus.org/tillingpages/homepage.htm). Identification of L. japonicus lines with mutations in CYP79D3 and CYP79D4 or in the as yet unknown transcription factor that regulates their expression might yield truly acyanogenic L. japonicus lines. This would provide useful tools to dissect unknown functions of cyanogenic glucosides and rhodiocyanosides in plant fitness and adaption to biotic and abiotic stresses. They may also approve valuable in unravelling the impact of cyanogenic glucosides and rhodiocyanosides in interactions with Zygaena larvae and plant development (Zagrobelny et al. 2004; Zagrobelny et al. 2007).
1.5 Expression of M. esculenta CYP79D2 in L. japonicus results in increased accumulation of linamarin but not lotaustralin
A surprising result of the expression of M. esculenta CYP79D2 in L. japonicus was therefore the accumulation of linamarin and lotaustralin but not rhodiocyanosides in roots (Fig. 7B). This demonstrates that roots could have an inherent capacity to synthesize cyanogenic glucosides but not rhodiocyanosides which strongly indicates the presence of two separate biosynthetic pathways for cyanogenic glucosides and rhodiocyanosides in roots and aerial tissues in L. japonicus in accordance with the differential expression of CYP79D3 and CYP79D4 (Forslund et al. 2004). The possibility that the root content of linamarin and lotaustralin was derived from translocation from the aerial parts to the roots cannot be ruled out. Unfortunately, the CYP71E1 orthologues encoding the putative cytochromes P450 that catalyzes the conversion of oxime into cyanohydrin in L. japonicus have not yet been identified and this hampers the understanding of how L. japonicus organizes the biosynthesis of cyanogenic- and cyanoalkenyl glucosides in aerial tissues; either as parallel metabolons or as a single promiscuous metabolon that synthesizes both sets of glucosides simultaneously.
When Z. filipendulae larvae were reared on L. japonicus wild type and L. japonicus 35S::CYP79D2, an increase in linamarin to lotaustralin ratio was observed in the larvae fed on L. japonicus 35S::CYP79D2 compared to wild type (Zagrobelny et al. 2007). These results demonstrate that the linamarin to lotaustralin ratio present in Zygaena larvae partly reflects the ratio in their dietary plants (Zagrobelny et al. 2007).
2 Concluding remarks
Predictive metabolic engineering of secondary metabolites is the key goal of many research programs. Yet in most cases, metabolic engineering of secondary metabolites is still in its infancy and metabolic engineering is subject to much trial and error. The lessons learned from metabolic engineering of cyanogenic glucosides may provide clues on how to proceed in other similar research initiatives and highlight important factors to be considered when plants are engineered with the purpose of obtaining altered profiles of their secondary metabolites.
The choice and number of different promoters that control expression of the transgenes appears to be critical, depending on the species. In some plant species several copies of the same promoter may be introduced without any adverse effects, while in other species this may be detrimental to the desired outcome of the experiment. This is exemplified by homozygous A. thaliana 3x plants that maintain transgene expression in the presence of ten copies of the CaMV 35S promoter and accumulate high levels of dhurrin (Tattersall et al. 2001). In contrast, heterozygous transgenic L. japonicus 3x plants harboring the same constructs but only half the copy number (this paper) are subject to gene silencing. At the other end of the scale is the consistent silencing of transgenes driven by the CaMV 35S promoter in gentian (Gentiana triflora × G. scabra) (Mishiba et al. 2005).
In the case of L. japonicus, application of the CYP79D3 promoter regulating endogenous cyanogenic glucoside biosynthesis (Forslund et al. 2004) would be an obvious alternative to CaMV 35S for driving expression of the three S. bicolor cDNAs that encode the enzymes involved in dhurrin biosynthesis. The risk of TGS due to the presence of several copies of the same promoter could be significantly reduced by expression of a multigene expression construct consisting of the genes encoding the three biosynthetic enzymes and the selective marker under control of one single promoter. By separation of the four proteins with the viral 2A peptide (El Amrani et al. 2004), the individual enzymes are predicted to be co-translationally cleaved to yield CYP79A1, CYP71E1, UGT85B1 and the selective marker protein. Alternatively, or in combination with the 2A polyprotein strategy, the use of an inducible promoter would allow selection of transgenic explants without the risk of counter selection for reduced expression of the transgenes. The use of a wound inducible promoter would add a new dimension to the study of cyanogenic glucosides and their impact on plant–insect interactions by changing their characteristics from being regarded as phytoanticipins (preformed defense compounds) to phytoalexins (defense compounds synthesized in response to herbivore or pathogen attack).
Sorghum bicolorCYP79A1 and CYP71E1 were transformed into A. thaliana, N. tabacum and L. japonicus with the anticipation that an endogenous UDP glucosyl transferase possessing broad substrate specificity (Hansen et al. 2003; Jones et al. 1999) would readily glucosylate the product p-hydroxymandelonitrile to yield dhurrin. This was not the case for any of these plants, which emphasizes the specificity of S. bicolor UGT85B1 and the need for insertion of the entire dhurrin biosynthetic pathway in the transformation experiments. The highly specific S. bicolor UGT85B1 glucosyl transferase fulfills the requirement for high substrate specificity combined with the ability to form a metabolon with the two cytochromes P450 in the pathway (Jones et al. 2000; Jørgensen et al. 2005b; Møller and Conn, 1980).
Cyanogenesis is the ability of a living organism to release toxic HCN. Cyanogenesis requires the presence, i.e. the ability of the living organism to synthesize the cyanogenic compound as well as the presence of enzymes that are able to cleave this compound with a concomitant release of HCN (Conn, 1980). In a previously non-cyanogenic plant, introduction of cyanogenesis would therefore typically require not only the introduction of the enzymes involved in biosynthesis but also the catabolic enzymes to facilitate a rapid HCN release. In addition, the cyanogenic compounds produced should be compartmentalized separately from the degrading enzymes until tissue disruption. Though β-glucosidases are present in A. thaliana plants, the lack of a specific dhurrin β-glucosidase in A. thaliana 3x compromises the utility of these transgenic plants for the study of plant–insect interactions because the cyanide release resulting from tissue damage is slow in comparison to that observed in naturally occurring cyanogenic plants. In order to fully exploit the HCN potential of the A. thaliana 3x lines, a cDNA encoding a β-glucosidase with high activity towards dhurrin is currently being introduced into A. thaliana 3x.
The major lesson learned from metabolic engineering of cyanogenic glucosides is that detailed knowledge of biosynthesis, regulation, transport, degradation and metabolic cross-talk is a prerequisite for performing predictive metabolic engineering. Even possessing this information, the facility of changing the metabolome of a given plant also depends on the choice and number of promoters in concert with the plant’s ability to successfully produce active heterologous enzymes and accommodate the biosynthetic product and possible toxic intermediates thereof.
We thank present and former members of the Cyanogenic Glucoside and Molecular Evolution group for their contributions to the work presented in this paper. Ms Susanne Jensen and Mrs Charlotte Sørensen are thanked for excellent technical assistance. We are very grateful to Mr Steen Malmmose for taking great care of the M. esculenta and L. japonicus plants. Financial support from the Danish National Research Foundation and a PhD stipend from University of Copenhagen to AVM are greatly acknowledged