Introduction

Terpenoids constitute the largest family of natural plant products with over 30,000 members possessing important biological and physiological functions (Sacchettini and Poulter 1997). Terpenoids are derived from geranyl diphosphate (GDP) which is synthesized by sequential head to tail addition of isopentenyl pyrophosphate and its allelic isomer dimethylallyl diphosphate (Wise and Croteau 1998). Synthesis of isopentenyl pyrophosphate and dimethylallyl diphosphate proceeds via cytosolic mevalonate (MVA) and plastid methylerythritol phosphate (MEP) pathways (Mahmoud and Croteau 2002; Hampel et al. 2006; Fig. 1). A cross-talk between these two pathways has also been reported (Hampel et al. 2005). Supply of GDP is critical in realizing the yield of terpenoids (Nogués et al. 2006), therefore, studies on regulation of genes in GDP biosynthesis assume central importance.

Fig. 1
figure 1

Schematic pathway for picrosides biosynthesis (adapted from Mahmoud and Croteau 2002). Geranyldiphosphate (GDP) can be derived from mevalonate (MVA) or methylerythritol phosphate (MEP) pathway. GDP yields iridoid, which is converted into picrosides in the presence of glucose and cinnamic acid or vanillic acid. Encircled numbers represent enzyme catalyzing the corresponding reaction step as follows: 1 1-deoxy-D-xylulose-5-phosphate synthase; 2 1-deoxy-D-xylulose-5-phosphate reductoisomerase; 3 2-C-methylerythritol 4-phosphate cytidyl transferase; 4 4-(cytidine-5′-diphospho)-2-C-methylerythritol kinase; 5 2-C-methylerythritol-2,4-cyclophosphate synthase; 6 1-hydroxy-2-methyl-2-(E)-butenyl-4-diphosphate synthase; 7 1-hydroxy-2-methyl-2-(E)-butenyl-4-diphosphate reductase; 8 acetoacetyl CoA thiolase; 9 3-hydroxy-3-methylglutaryl coenzyme A synthase; 10 3-hydroxy-3-methylglutaryl coenzyme A reductase; 11 mevalonate kinase; 12 phosphomevalonate kinase; 13 mevalonate-5-pyrophosphate decarboxylase; 14 isopentenyl pyrophosphate isomerase; 15 geranyldiphosphate synthase. Solid arrows indicate known steps, whereas dotted arrows indicate unknown steps

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Regulation of gene expression at the level of transcription is a major control point in many biological processes. Transcriptional regulation is achieved through binding of transcription factors to short consensus sequences of DNA known as cis-acting regulatory elements or motifs, usually located in the promoter regions, upstream of the coding sequences (Udvardi et al. 2007). It is generally believed that genes having similar expression patterns contain common motifs in their promoter (Klok et al. 2002). Thus, a common set of transcription factors co-regulate expression of genes and hence the pathway.

Picrosides, the medicinally important hepatoprotectants (Floersheim et al. 1990), are synthesized by picrorhiza (Picrorhiza kurrooa Royle ex Benth.), an endangered plant species of family Scrophulariaceae. The species is distributed between 3,000-5,000 m above mean sea level in Himalaya (Chettri et al. 2005). Indiscriminate and extensive harvesting and lack of organized cultivation of the plant has threatened its status in wild and listed as ‘endangered’ species by International Union for Conservation of Nature and Natural Resources (Nayar and Sastri 1990).

Picrosides are terpenoids with an iridane skeleton of monoterpene origin (Fig. 1). Previous studies have shown that 3-hydroxy-3-methylglutaryl coenzyme A reductase (hmgr) and 1-deoxy-D-xylulose-5-phosphate synthase (dxs) are regulatory genes in terpenoids biosynthesis (Korth et al. 2000; Hsieh and Goodman 2005) for which no information is available in picrorhiza. In the present work, we cloned these genes from picrorhiza, analyzed the cis-acting elements, and the motifs for light and temperature regulation were studied. Electrophoretic mobility shift assay (EMSA) confirmed binding of proteins to the identified motifs. Gene expression analysis supported the results on transcriptional regulation of hmgr and dxs by light and temperature. A positive relationship between expression of these genes and picrosides content was obtained.

Materials and methods

Plant material

Plants of picrorhiza (P. kurrooa Royle ex Benth.) were collected from its natural habitat at Rohtang pass (4,000 m altitude, 32°23′ N, 77°15′ E, India) and brought to the Institute at Palampur (1,300 m altitude; 32°06′ N, 76°33′ E, India). These were transplanted in plastic pots (20 cm height × 20 cm top diameter × 12 cm bottom diameter) containing soil, sand, and farm yard manure mixture in a ratio of 2:1:1, and were maintained in the experimental farm of the Institute. Plants were allowed to acclimatize for 3 months before start of the experiment. Experiments were performed on fifth leaf (position with respect to the top apical leaf designated as first leaf), which was harvested at the designated time, followed by freezing in liquid nitrogen and storage at −80°C for further use.

Raising of in vitro shoot cultures of picrorhiza

Young leaves of picrorhiza were collected from plants grown in the experimental farm, washed with tween-20, and surface-disinfected with solution containing 0.05% (w/v) streptomycin sulfate and 0.05% (w/v) bavistin for 20 min. The explants were further washed with distilled water and surface sterilized with sequential treatment of 70% ethanol for 45 s and mercuric chloride 0.05% (w/v) for 10 min. Subsequently, these were washed with autoclaved distilled water and inoculated on Murashige and Skoog (MS) medium (Murashige and Skoog 1962) supplemented with 3% sucrose. Prior to autoclaving, pH of the medium was adjusted to 5.7 followed by addition of 0.8% agar. Unless indicated, regenerated plantlets were incubated at 25 ± 2°C with 16 h photoperiod (photosynthetic photon flux density, 70 µmol m−2 s−1).

Exposure of picrorhiza to temperature and light

Potted plants were shifted from the experimental farm to the plant growth chamber maintained at 25 ± 2°C (16 h photoperiod; photosynthetic photon flux density, 350 µmol m−2 s−1). After 5 days, a few pots were shifted to another growth chamber maintained at 15 ± 2°C. Data was collected on day 12th after shifting as detailed in the figure legends.

Our data on the effect of temperature showed poor expression of the targeted genes at 25°C, which led us to study the effect of light in plants maintained at 15°C. For dark treatment, plants were covered with a cardboard box to restrict entry of light and the samples were harvested at 12 h from start of the treatment. Any long-term experiment with the plants exposed to dark could lead to possible starvation thus limiting the carbon pool. In order to differentiate the impact of light through effect on carbon pool or the light per se, plants were raised in vitro through tissue culture as detailed above. Four-week-old plantlets raised at 25 ± 2°C were transferred to test tubes containing MS medium supplemented with 3% sucrose and maintained at 15 ± 2°C in light and complete darkness, separately. Samples were harvested on day 6 of the treatment for analysis of gene expression and picrosides content. Beyond day 6, plants kept in dark showed etiolation and yellowing.

Cloning of cDNAs of pkhmgr and pkdxs

RNA was isolated from leaf tissue using the method of Ghawana et al. (2007) and digested with DNase 1 (RNase-free) using Message Clean® Kit (GenHunter® Corporation, USA). Complementary DNA (cDNA) was synthesized as described by Singh et al. (2004). Degenerate primers for hmgr and dxs were designed from the conserved regions of corresponding genes reported for different plant sources and the partial gene sequences were amplified by PCR as detailed in Table 1. The amplicons were cloned in pGEM-T Easy Vector (Promega, USA), plasmids were isolated using Qiagen Plasmid Mini-isolation Kit (Qiagen, GmbH), and sequencing was performed using BigDye terminator cycle sequencing mix (Version 3.1; Applied Biosystems, USA) using an automated DNA sequencer (ABI PRISM™ 310 and 3130 xl Genetic Analyzer, Applied Biosystems, USA). Protocols were followed essentially as described by the respective manufacturers.

Table 1 Oligonucleotide sequences and PCR conditions used in the present work
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Full-length cDNAs were cloned by performing rapid amplification of cDNA ends (RACE; SMART™ RACE cDNA Amplification Kit; Clontech, USA) as per the manufacturer’s instructions using the gene specific primers (hmgr5′R, hmgr3′R, dxs5′R and dxs3′R; Table 1). These primers were designed based upon the partial sequences of the genes as cloned above. After aligning the sequences obtained by 5′ and 3′ RACE, full-length cDNA was amplified using the end sequences, cloned in pGEM-T Easy Vector (Promega, USA) and confirmed by sequencing.

Wherever needed, cDNA-sequences were analyzed using BLASTN, BLASTX, and BLASTP programs of National Center for Biotechnology Information (NCBI) with default parameters (http://www.ncbi.nlm.nih.gov/). Secondary structure of the deduced amino acid sequence was analyzed using Self-Optimized Prediction Method with Alignment (SOPMA; http://www.npsa-pbil.ibcp.fr/).

Construction of genome-walking libraries and cloning of promoters

DNA was isolated from leaf tissue using cetyl-trimethylammonium bromide-based procedure (Doyle and Doyle 1987), treated with RNase (DNase-free) and purified with phenol/choloform for constructing libraries for genome walking (GenomeWalker™ Universal Kit; BD Bioscience, Clontech, USA). Genomic DNA was digested for 16 h with restriction enzymes DraI, EcoRV, PvuII, and StuI, separately to create blunt-end fragments and purified using phenol/chloroform. The restricted fragments were ligated to adaptors supplied by the manufacturer to produce four separate libraries. Promoter sequences were amplified using the Advantage Genomic Polymerase Mix (BD Bioscience, Clontech, USA) in a thermocycler (i-cycler, Bio-Rad, USA). Primary PCR was performed with gene-specific primers (hmgrR-p and dxsR-p; Table 1) and the outer adaptor primer (AP1, supplied by the manufacturer) using all the four genomic libraries, followed by secondary PCR with nested gene-specific primers (hmgrR-s, dxsR-s; Table 1) and the nested adaptor primer (AP2, supplied by the manufacturer). The PCR fragments were analyzed on a 1.2% agarose gel. Amplicons were purified using QIAEX®II Gel Extraction Kit (Qiagen, GmbH), cloned into pGEM-T Easy Vector (Promega, USA), and sequenced. Sequence information was also used for subsequent rounds of genome walking (if needed).

Sequences were analyzed in silico for detection of various motifs using “plant cis-acting regulatory DNA elements” (PLACE) database (http://www.dna.affrc.go.jp/PLACE ; Higo et al. 1999). PLACE has advantage over the other eukaryotic databases such as TFD and TRANSFAC in terms of containing larger numbers of eukaryotic cis-elements from vascular plants (Higo et al. 1999). Multiple Expectation-Maximization for Motif Elicitation was used for the detection of common motifs in both the promoters with motif width set to 6-9 and the number of motifs to be detected was set to 10 (http://meme.sdsc.edu/meme/meme). Since varying length of promoters was obtained for the two genes, the analysis of plus-strands was confined to 1,000 nucleotides upstream to the translation start site.

Reverse transcriptase-PCR analysis

DNA-free RNA was used to synthesize cDNA using Accuscript™ High Fidelity Ist strand cDNA Synthesis Kit (Stratagene, USA). This cDNA was to be used as template for Reverse transcriptase-PCR reaction using gene specific primers (hmgrF, hmgrR; dxsF, dxsR) as mentioned in Table 1. Cycling conditions were optimized to obtain amplification under the exponential phase. 26S rRNA based primer pair was used as internal control for expression studies (Singh et al. 2004). Amplicons were analyzed and quantified using the Alpha DigiDoc Gel Documentation and Image analysis system (Alpha Innotech, USA). Each experiment was repeated at least twice with three biological replicates each time and the representative figure of one experiment is shown in the manuscript.

Extraction and estimation of picrosides

Picrosides were estimated as described by Dutt et al. (2004). Leaf tissue was harvested, washed with distilled water, blotted dry, weighed, then frozen in liquid nitrogen and stored at −80°C. The frozen samples (20 mg) were ground to fine powder in liquid nitrogen using pestle and mortar followed by addition of 1.0 ml of 80% methanol with intermittent grinding for 1 min. Extract was transferred to a centrifuge tube and the pestle and mortar was rinsed with 1.0 ml of 80% methanol to recover the leftover sample. Extracts were pooled, centrifuged at 15,000×g for 15 min and the supernatant was used for picrosides estimation. Samples were filtered through 0.45 micron filter (Millipore, USA) for high pressure liquid chromatography analysis using LC 4000 module, 2487 dual λ absorbance detector (both from Waters, UK), and LiChrosorb® RP-18, 250 X 4.0 mm (Hibar) column (Merck & Co., Inc., USA). The mobile phase (flow rate, 1.0 ml/min) consisted of a mixture of trifluoroacetic acid (0.05%) and methanol-acetonitrile (1:1) in 70:30 ratio. Picrosides were monitored at 270 nm and quantified using pure picroside-I (P-I) and picroside-II (P-II) as standards.

Nuclear proteins extraction and electrophoretic mobility shift assay

Leaf tissue (20 g) was harvested from the plants growing at temperature of 15°C and a photoperiod of 16 h, and ground in pre-chilled pestle-mortar using extraction buffer [10 mM HEPES (pH 7.8), 10 mM KCl, 10 mM MgCl2, 5 mM EDTA, 1 mM DTT, 250 mM Sucrose, 0.2 mM PMSF and 0.5% Triton X-100] to isolate nuclear proteins essentially as described by Busk and Pages (1997). Oligonucleotides (30 mer) possessing low temperature responsive element (LTRE; CCGAA) and light responsive element (GATA), as present in promoters of pkhmgr and pkdxs, were designed having sequences: 5′-GAATTCAAAACTTTAACTGAAACCGAAAAATTGAGAAA-3′ (LTRE-1); 5′-GAATTCTTTCTCAATTTTTCGGTTTCAGTTAAAGTTTT-3′ (LTRE-2; complementary to LTRE-1); 5′-GTGCCAGTGTTGATATTATAAGTGCCAGTG-3′ (GATA-1); 5′-CACTGGCACTTATAATATCAACACTGGCAC-3′ (GATA-2; complementary to GATA-1). Equal moles of complementary oligonucleotides were mixed and annealed by incubating at 95°C for 5 min followed by slow cooling to room temperature. For preparation of probe for EMSA (Hennighausen and Lubon 1985), double-stranded oligonucleotides were radio-labeled with α-32P-dATP (3,000 Ci/mmol) using Klenow DNA polymerase I enzyme (USB, USA) that filled up the recessive ends. Recessive ends were created by incorporating GAATTC at 5′ end of the oligonucleotides. Binding of protein was carried out in 1× binding buffer [30 mM HEPES (pH 7.8), 60 mM KCl, 0.3 mM EDTA, 0.5 mM DTT and 10% glycerol] containing 4 µg of nuclear protein. For specific competition, unlabeled competitor DNA was added in excess to the binding mixture. After 10 min incubation at room temperature, probe (10,000 cpm) was added and incubated further for 20 min. Samples were loaded onto a 6% non-denaturing polyacrylamide gel and run in 0.5× TBE buffer [1× TBE is 50 mM Tris-borate (pH 8.2), 1 mM EDTA] at 15 V/cm. Gel was blotted onto 3MM filter paper (Whatman, Clifton, NJ), dried under vacuum at 80°C for 2 h, and subjected to autoradiography.

Results and discussion

Hepatoprotective activity of picrorhiza owes to the presence of monoterpenoid picrosides (P-I and P-II; Luper 1998). Since terpenoids are derived from GDP that can be synthesized both from cytoplasmic MVA and plastidic MEP pathways (Croteau et al. 2005), it is important to study the regulation of these two pathways as feeders of GDP. Enzyme 3-hydroxy-3-methylglutaryl coenzyme A reductase (encoded by hmgr) catalyzes reduction of 3-hydroxy-3-methylglutaryl coenzyme A into mevalonate and is regarded as rate-limiting step in the MVA pathway (Wang et al. 2007), for example in potato (Solanum tuberosum) and Arabidopsis (Yang et al. 1991; Leivar et al. 2005). Enzyme 1-deoxy-D-xylulose-5-phosphate synthase (encoded by dxs) catalyzes the first rate-limiting step involving condensation of pyruvate and glyceraldehyde-3-phosphate to produce 1-deoxy-D-xylulose-5-phosphate in the MEP pathway. The gene has been found to be regulatory (Muñoz-Bertomeu et al. 2006; Estevez et al. 2001), such as in tomato (Lycopersicon esculentum), periwinkle (Catharanthus roseus) and Arabidopsis (Lois et al. 2000; Chahed et al. 2000; Juan et al. 2001). Since hmgr and dxs are regarded as important genes of terpenoid biosynthesis pathway (Fig. 1), the present work studied these genes, their promoters, and the picrosides content to identify the gene regulation and its possible relationship with the picrosides content in picrorhiza.

Degenerate primers (Table 1) followed by RACE led to amplification of 2,245 bp and 2,317 bp of pkhmgr (accession no. DQ347962) and pkdxs (accession no. EU561005), respectively. The cDNAs consisted of 106 and 50 bp of 5′ untranslated region; 453 bp, and 203 bp of 3′ untranslated regions; and open reading frames of 1,686 and 2,064 bp for pkhmgr and pkdxs, respectively. Deduced proteins (hereinafter referred to as pkHMGR and pkDXS, respectively) had molecular masses/isoelectric points of 60.65 kDa/7.24 and 73.63 kDa/8.60, respectively. BLASTX analysis showed extensive identity (>70%) of the deduced proteins with those from other plant sources (Electronic Supplementary Fig. S1).

In silico analysis showed the presence of corresponding conserved domains in both of the deduced proteins (Electronic Supplementary Fig. S2). HMG coenzyme A reductase class I domain was found to be present between amino acid (aa) positions 162 and 551 in pkHMGR. In pkDXS, three characteristic domains were recorded. These were: thiamine pyrophosphate binding domain (between aa positions 77 and 334), transketolase pyrimidine binding domain (between aa positions 364 and 527), and transketolase–c terminal domain (between aa positions 545 and 667).

SOPMA analysis revealed 46.52% α-helices, 4.99% β-turns, 14.62% extended strands and 33.87% random coils in pkHMGR. In pkDXS, α-helices, β-turns, extended strands, and random coils were 36.97%, 6.84%, 14.56%, 41.63%, respectively (Electronic Supplementary Table S1, Fig. S3). Deduced secondary structures were in agreement with those reported for functional genes in other plant systems (Electronic Supplementary Table S1). Various in silico analyses suggested the cloned genes to be functional as has been reported by the other studies (Walter et al. 2002; Wang et al. 2007).

Cloning and analysis of promoters of pkhmgr and pkdxs

The promoters of pkhmgr (Upkhmgr, 1,065 bp; accession no. FJ228691) and pkdxs (Upkdxs,1,146 bp; accession no. FJ228692) were cloned by genome walking. In silico analysis was limited to 1 kb sequences upstream of translation start site for consistency in comparative analysis of the two promoters under study. Table 2 lists various motifs along with their function and location in the two promoters analyzed. A total of 73 types of motifs were identified in Upkhmgr and Upkdxs, out of which 24 motifs were common to both, whereas 27 and 22 motifs were specific to Upkhmgr and Upkdxs, respectively (Table 2). In Upkhmgr, number of motifs identified for basal transcription, light responsiveness, low temperature responsiveness, tissue specificity (leaf, seed, root, flower, meristems), hormone (gibberellins, cytokinins, abscisic acid, and auxin) responsiveness and biotic factors were 11, 17, 7, 61, 19, and 11, respectively. These figures for Upkdxs were 9, 16, 8, 28, 23, and 7 for the above motifs in the same order.

Table 2 Location of motifs with respect to the translation start site detected in Upkhmgr and Upkdxs using PLACE database (http://www.dna.affrc.go.jp/PLACE). Motifs at serial number 25-73 are specific to either Upkhmgr or Upkdxs
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Ecological niche of picrorhiza is characterized by an environment of low temperature and high light intensity (Chandra 2004). Light responsive motifs in both the promoters were spread throughout the promoter (Fig. 2) and these include: GATA box (WGATAR; Gilmartin et al. 1990), GT-1 (GRWAAW; Villain et al. 1996), TATA box (TTATTT; Tjaden et al. 1995), and I box (GATAA; Terzaghi and Cashmore 1995). GATA factors were first identified as proteins that interact with conserved WGATAR (W = T or A; R = G or A) motifs involved in erythroid-specific gene expression in vertebrates (Evans et al. 1988) and have been mostly implicated in light-dependent gene regulation in plants (Reyes et al. 2004). In Arabidopsis, Teakle et al. (2002) established that different families of GATA-binding factors bind to DNA sequence elements containing GATA or GAT sequence. It was suggested that GATA core sequence in the GATA motif was important for binding relevant transcription factors and the flanking sequences are non-consequential.

Fig. 2
figure 2

Nucleotide sequences of Upkhmgr and Upkdxs promoters cloned from picrorhiza. One thousand nucleotides upstream to translation start site were analyzed. The nucleotide just upstream to the first nucleotide of translation start codon is numbered as −1 and subsequent upstream numbering is in descending order. Light and temperature responsive motifs are boxed and underlined, respectively

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GT-1 box is a regulatory motif usually found in tandem repeats in the promoter region of many different plant genes regulated by light (Villain et al. 1996). Apart from light, GT-1 is also shown to be regulating salt and pathogen induced gene expression (Park et al. 2004) and hence is not considered very specific in relation to regulation by light.

Apart from the above motifs common in the two promoters, promoter-specific light responsive motifs were also detected. In Upkhmgr, GCCAC (SORLIP) box was identified which is found to be over-represented in the phytochrome A-induced promoters (Hudson and Quail 2003). Upkdxs possessed CGGATA (REbeta) box, which is shown to be required for phytochrome regulation in Lemna gibba Lhcb21 gene promoter (Degenhardt and Tobin 1996). Yet another light responsive box GGTTAA was identified in Upkdxs, which has been reported to be present in upstream sequences of the light-responsive pea rbcS-3A gene (Green et al. 1988).

Among the various light responsive motifs analyzed in silico, GATA core sequence was specific to light and was present in Upkhmgr and Upkdxs. Therefore, GATA core was selected to study its interaction with putative transcription factors by EMSA. Results showed prominent shifts for GATA motif (Fig. 3 lane 5) suggesting binding of putative transcription factors to the motif in vitro.

Fig. 3
figure 3

Electrophoretic mobility shift assay showing DNA-protein interaction. Lane 1 radio-labeled low temperature responsive sequence (LTRE; 5′-GAATTCAAAACTTTAACTGAAACCGAAAAATTGAGAAA-3′); lane 2 radio-labeled LTRE sequence incubated with nuclear protein; lane 3 radio-labeled LTRE sequence incubated with nuclear protein and unlabeled (functioning as specific competitor) LTRE sequence; lane 4 radio-labeled light responsive (GATA) sequence (5′-GTGCCAGTGTTGATATTATAAGTGCCAGTG-3′); lane 5 radio-labeled GATA sequence incubated with nuclear protein; and lane 6 radio-labeled GATA sequence incubated with nuclear protein and unlabeled (functioning as specific competitor) GATA sequence. Core LTRE and GATA sequences are underlined

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Low temperature responsive motifs were investigated to examine various cis-acting regulatory elements and trans-acting factors that have been reported to be involved in plants responses to temperature. Presence of several motifs involved in low temperature responses were detected in both the promoters analyzed (Table 2, Fig. 2). These included low temperature responsive elements (LTRE, CCGAC/CCGAA) and dehydration responsive elements [DREs: CANNTG (MYC), WAACCA, CNGTTR, TAACTG, YAACKG (MYB), ACGT, and MACGYGB (ABRE)]. LTREs containing pentanucleotide CCGAC/CCGAA have been identified in a number of promoters of a low-temperature-responsive gene such as in Brassica napus (BN115) and Hordeum vulgare (ABA-regulated barley gene, HVA1; Dunn et al. 1998). EMSA showed the importance of CCGA core sequence within the pentanucleotide LTRE for DNA protein interaction. Motifs related to water stress/drought viz; MYC, MYB, and ABRE like boxes are involved in ABA-mediated responses to various adverse environmental cues including cold (Agarwal et al. 2006; Choi et al. 2000).

LTRE core sequence CCGA was present in Upkhmgr and Upkdxs and was used in DNA-protein interaction studies. Results showed prominent shifts for CCGA motif (Fig. 3 lane 2) suggesting binding of putative transcription factors to the motif in vitro.

Data on in silico analysis and EMSA suggested the possible functionality of light and temperature responsive motifs in promoters of both the genes. The next logical question was if these genes are also regulated by light and temperature? And how does it correlate with the picrosides content under these conditions.

Effect of light and temperature on expression of pkhmgr and pkdxs, and picrosides content

Expression of pkhmgr and pkdxs was higher by 216% and 286%, respectively, at 15°C as compared to those at 25°C (Fig. 4a, b). Also, total picrosides increased by 22% at 15°C as compared to the plants maintained at 25°C (Fig. 4c). Since expression of pkhmgr and pkdxs was very poor at 25°C, experiment on the effect of dark was performed at 15°C. In a span of 12 h, expression of pkhmgr and pkdxs was higher by 70% and 112% in light as compared to that observed under dark (Fig. 5a, b). Picrosides content in light was higher by 163% of that observed under dark conditions (Fig. 5c).

Fig. 4
figure 4

Effect of temperature on expression of pkhmgr and pkdxs, and picrosides content in fifth leaf of picrorhiza. Panel a shows gene expression, wherein 26S rRNA was used as a marker for equal loading. Panel b shows integrated density value (IDV) of each amplicon as obtained in panel a. Panel c shows change in picrosides content. Values in panel b and c are average of three separate biological replicates; error bar represents standard error of the mean

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Fig. 5
figure 5

Effects of light and dark on expression of pkhmgr and pkdxs, and picrosides content in fifth leaf of picrorhiza plants (a, b, c) and plantlets raised through tissue culture (d, e, f). Panels a and d show gene expression, wherein 26S rRNA was used as a marker for equal loading. Panels b and e show integrated density value (IDV) of each amplicon as obtained in panels a and d. Panels c and f show changes in picrosides content. Values in panel b, c, e, and f are average of three separate biological replicates; error bar represents standard error of the mean

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Dark conditions might starve the plants and it is likely that gene expression and picrosides content in dark could be a reflection of the effect of carbon limitation. Light would affect carbon pool through photosynthesis and the role of carbon pool in regulating secondary metabolites has been shown in Hypericum perforatum L. (Mosaleeyanon et al. 2005). Also, the possibility exists that light modulated gene expression independent of carbon pool (Fey et al. 2005). Hence, the experiment was carried out using in vitro raised plantlets with the medium fortified with sucrose (3%). In this system also, expression of pkhmgr and pkdxs was higher by 173% and 112% in light as compared to that recorded under dark (Fig. 5d, e). Also, the picrosides content in light was higher by 389% as compared to those in dark (Fig. 5f). Gene expression was in agreement with the promoter data wherein the motifs for light (e.g., GATA) and low temperature (e.g., LTRE) were present. EMSA also showed binding of the putative transcription factors to these motifs (Fig. 3).

The role of light and temperature in modulating a range of terpenoids and the corresponding transcripts has been documented, but there is no universal behavior and it varies depending upon the metabolite under study and the plant species. The same gene, hmgr for example, is stimulated by light in Triticum aestivum (Aoyagi et al. 1993), pea (Wong et al. 1982) and potato (Korth et al. 2000) but is downregulated by light in Lithospermum erythrorhizon (Lange et al. 1998). Effect of light on the activity of hmgr promoter has been reported that explained the light mediated alteration in hmgr transcripts (Learned and Connolly 1997). Gene dxs has also been reported to be regulated by light in Arabidopsis (Hsieh and Goodman 2005; Cordoba et al. 2009).

Soitamo et al. (2008) showed that light at low temperature induced expression of genes involved in synthesis of phenylpropanoids, carotenoids, and terpenoids. In Arabidopsis, light-dependent flavonoid (Fuglevand et al. 1996) and phenylpropanoid (Hemm et al. 2004) biosynthesis has been attributed to: (1) upregulation of relevant genes at transcriptional level, and (2) the involvement of the primary photoreceptors- phytochrome B and cryptochrome. Light-mediated accumulation of reducing equivalents in the stroma and reactive oxygen species has been reported to function as signals from chloroplasts to the nucleus leading to altered gene expression (Fey et al. 2005). Low temperature mediated increase in secondary metabolites in Arabidopsis has mainly been attributed to the transcriptional upregulation of genes of secondary metabolism, which in turn, has been suggested due to the over-expression of relevant transcription factors at low temperature (Hannah et al. 2005). It is also possible that upregulation of the two regulatory genes of the picrosides biosynthesis pathway by light and low temperature increased carbon partitioning towards terpenoid metabolism resulting in higher picrosides content in picrorhiza.

Our results that light and low temperature favor picrosides accumulation are in agreement with the picrosides levels in natural population of picrorhiza, wherein it has been reported that picrosides content increased by 135% in the plants growing at high (4,145 m) as compared to the low (1,350 m) altitude (Singh et al. 2005). Increase in altitude accompanies decrease in temperature and increase in light quanta (Streb et al. 1998). Molecular data on promoter analysis, gene expression, and picrosides content validated the importance of light and temperature in regulating picrosides content in picrorhiza.